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This is to certify that the

dissertation entitled

The Metabolism of D-Lactate and Structurally
Related Organic Acids in Chlamydomonas Reinhardtii

 

presented by

Diane White Husic .

has been accepted towards fulfillment
of the requirements for

Ph . D . degree inBiochemis try

77. EW

Major professor

Date_.1nne 19; 1986

MS U is an malaria: Aca’on/Equal Opportunity Institution 0-12771

 

MSU

 

 

 

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'11 0 3 DJ

 

 

'IHE METAmLISM OF D-LAL'I‘ATE AND MY
RELATED WC ACIDS IN W REIMMRDTII

Diane White Husic

A DISSERTATICN

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

WOFPHIUBOHIY

Department of Biochemistry

1986

ABSTRACT

THE METABOLISM‘OF D—LACTATE AND STRUCTURALLY
RELATED ORGANIC ACIDS IN CHLAMYDOMONAS.REINHARDTII

By

Diane White Husic

During the initial minutes of anaerobiosis, 14C-labeled D—lactate,
derived from the photosynthetic sugar phosphate pool , accumlated in the
unicellular green alga, ailanydamonas reinhardtii. The production of the
D—isomer of lactate by algae is in contrast to plant and mnmalian cells
in which L—lactate is fanned. After initial lactate formation,
Chlamydomonas exhibits a mixed-acid type fermentation , thereby avoiding
lactate accumlation and enabling the cells to tolerate extended periods
of anaerobiosis.

A pyruvate reductase which catalyzes the formation of D—lsctate in
Chlamydomonas was partially purified and characterized. Lactate
produced anaerobically was metabolized only when Chlamydamonas cells were
returned to aerobic conditions, and reoxidation of the D-lactate was
apparently catalyzed by a mitochondrial membrane-bound.dehydrogenase,
rather than by the soluble py'ruvate reductase. Mutants of Chlamydamonas,

deficient in mitochondrial respiration, were used to demonstrate that

lactate metabolism was linked to the mitochondrial electron transport
chain. In addition, the oxidation of glycolate, a structural analog of
lactate, was also linked to mitochondrial electron transport in vivo.

The pyruvate reductase of Chlamydcmonas also catalyzed the reduction
of hydroxypyruvate and glycxylate, both of which are intermediates of the
oxidative photosynthetic carbon (CZ) cycle. Two other enzymes which
catalyze the reduction of hydroxypyruvate and glyoxylate, but not
pyruvate, were partially purified and characterized from Chlamydanonas
reinhandtii. These enzymes, NAbflzhydrmcypyruvate reductase and
NADpflzglyoxylate reductase, resembled the analogous C cycle enzymes from

2
higher plants .

am loving husband
for all the understanding, encouraganent and helpful

suggestions and criticism he provides

and
to all those at the Department of Chemistry at Northern Michigan

University who inspired my interest in the field of science

ii

ACKNOWLEDGEMENTS

I wish to thank Dr. Ed.Tolbert for providing me with the opportunity
and financial support to work in his laboratory and to follow a project
quite diverse from the research being pursued in his lab. Thanks also
for always finding a way to send me to all those scientific meetings
which provided valuable experience in presenting and.defending my data,
and the chance to meet numerous researchers in the field of plant
science.

I wish to thank those faculty members who served.on my graduate
committee: Dr. Shelagh FergusonrMiller, Dr. William wells, Dr. Charles
Sweeley, and Dr. Shauna Somerville. Special thanks to Dr. Clarence
Suelter, my "substitute" coulnittee manber, who filled in for both my mini-
masters defense and my final thesis defense. Additional thanks are in
order for Dr. Suelter for all of our discussions on enzyme kinetics and
purification tricks.

I am grateful to Dr. Elizabeth Harris at the Chlamydcmonas Culture
Center at Duke University who provided the mutants of Chlamydcmonas which
were deficient in mitochondrial respiration. These mutants were
essential for the experiments described in Chapter IV. Additional thanks

to Dr. R. K. Togasaki, Indiana University, Dr. B. Sears, Michigan State

iii

University, Dr. A. Wang, University of Iowa, Dr. M. Spalding, Iowa State
University, and Dr. S. Miyachi, University of Tokyo for the algal strains
which they provided.

I would also like to thank the Department of Biochemistry, the
National Science Foundation, and the McKnight Foundation for providing

financial support .

iv

TABLE OF CONTENTS
Page

LIST OF TABLES. . . . . . . . . . . . . . . . . . . . . . . . . . . . ix
LIST OF FIGURES . . . . . . . . . . . . . . . . . . . . . . . . . . . xi
LIST OF ABBREVIATIONS . . . . . . . . . . . . . . . . . . . . . . . .xiii
CHAPTER I. LITERATURE REVIEW . . . . . . . . . . . . . . . . . . . . 1
A. Anaerobic Metabolism . . . . . . . . . . . . . . . . . . . . 1
B. Anaerobic Metabolism in Chlamydbmcnas. . . . . . . . . . . . 7

C. D—Lactate Dehydrogenases . . . . . . . . . . . . . . . . . . 11
unicellular Green Algae . . . . . . . . . . . . . . . . . . 11
Yeast and Fungi . . . . . . . . . . . . . . . . . . . . . . 11
Bacteria. . . . . . . . . . . . . . . . . . . . . . . . . . 17
Animals . . . . . . . . . . . . . . . . . . . . . . . . . . 17
Evolutionary Significance . . . . . . . . . . . . . . . . . 18

D. The Oxidative Photosynthetic Carbon (C2) Cycle . . . . . . . 20
CHAPTER II. D—LACI'ATE ROWATION IN W REIMMRDTII . . . . 27
Introduction . . . . . . . . . . . . ... . . . . . . . . . . 28
Materials and Methods. . . . . . . . . . . . . . . . . . . . 30

Materials. . . . . . . . . . . . . . . . . . . . . . . . 30
Ciowth and Harvesting of Algae . . . . . . . . . . . . . 3O
C-Labeling of Algae. . . . . . . . . . . . . . . . . 31
Two-Dimensional Paper Chroma graphy . . . . . . . . . . 32
Analysis of the Products of m-Fixation by
ChlamydOmcnas in Decreasing 5 Concentrations. . . . 33
Determination of the Stereospecific Configuration of
the Lactate Produced by Chlamydcmonas reinhardtii. . 34
Formation of D—Lactate by the Pyruvate Reductase of
Chlamydcmonas. . . . . . . . . . . . . . . . . . . . 35
Estimates of Intracellular pH. . . . . . . . . . . . . . 36

V

Results and Discussion . . . . . . . . . . . . . . .

Anaerobic Formation of Lactate . . . . . . . . . . . .

Determination of the Configuration of the Lactate
Produced by Chlamydbmonas. . . . . . . . . . . . .

Effect of Lactate Accumulation on the Intracellular
pH of Chlamydbmonas reinhardtii. . . . . . . . . . .

CHAPTER III. PARTIAL PURIFICATION AND CHARACTERIZATION OF A
PYRUVATE REDUCTASE (DbLACTATE DEHYDROGENASE) FROM CHLAMYDOMOMAS

.REINHARDTTI .

Introduction . . .

Materials and Methods. . . . . .

mterials O O O I O O O O O O O 0
Preparation of Algal Extracts. .

Enzyme Assays. . . . . . . . . . . .

49

50

51

51

51
52

Partial Purification of the Pyruvate Reductase from
Chlamydbmonas. . . . . . . . . . . . . . . . . . .

ksults O O O O O O O O O O O O O 0 O O O O O O O O 0 O O 0

Partial Purification of the Pyruvate Reductase
from Chlamwdbmonas. . . . . . .
Properties of the Partially Purified.Pyruvate
Reductase from Chlamwdbmonas. .
Effect of Phosphate and.0ther Anions on Pyruvate
Reductase . . . . . . . . . . . . . . . . . . .
Inhibition of the Pyruvate Reductase by various

Compounds . . . . . . . . . . . . . . . . .
Evidence for the Constitutive Nature of Pyruvate

Reductase . .

Discussion .

vi

79

81

CHAPTER IV. INHIBITION OF D-LACTATE AND GLYCOLATE METABOLISM IN A
MUTANT OF CIRAMflQXIIWWQSJRELNHARDTTI'DEFICIENT IN MITOCHONDRIAL
WMTIW C O O O O O O O O O O O O O O O O O O O O O O O O O

I "tram t i on O O O O O O O O O O O O O O O O O O O I O I O 0

Materials and Methods. . . . . . . . . . . . . . . . . . .

Materials. . . . . . . . . . . . . . . . . . . . . . .
Preparation of Algal Cell Suspensions and Crude
Homogenates. . . . . . . . . . . . . . . . . . . .
says . . . . . . . . . . . . . . . . . . . . . . . .
C-Labeling and Chase Experiments . . . . . . . . . .
Analysis of C-Labeled Products of Chlamydbmonas. . .

ksults O O O O O O O O O O O O O O O O O O O O O O O O O

D-Lactate Metabolism in Chlamydbmonas. . . . . . . .
Evidence that the Glycolate Dehydrogenase is Linked
to the Mitochondrial Electron Transport. . . . .
Evidence that DbLactate Metabolism is Linked to
Mitochondrial Electron Transport . . . . . . .
Evidence for the Persistence of Mitochondrial
Respiration in the Light During Photosynthesis
in Chlamydamanas . . . . . . . . . . . . . . . .

DiSCUSSion O 0 O O O O O O O O O O O O 0 O I O 0

CHAPTER V. PROPERTIES OF NADflzHYDROXYPYRUVATE REDUCTASE AND
NADPH:GLXOKYLATE REDUCTASE IN ALGAE. PARTIAL PURIFICATION AND
CHARACTERIZATION FROM CHLAMYDOMONAS.REINHARDTTI . . . . . . .

I ntrwm t i on O O O O O O O O O 0 O O O O O O O O 0 O O O 0

Materials and Methods. . . . . . . . . . . . . . . . . . .

Materials. . . . . . . . . . . . . . . . . . . . . .
Preparation of Algal Crude Hemogenates . . . . . . . .
Enzyme Assays. . . . . . . . . . . .
Partial Purification of the Hydroxypyruvate and
Glyoxylate Reductases from Chlamydomonas . . . .
Mblecular Weight Determination of the NADflzHydroxy-

pyruvate Reductase . . . . . . . . . . . .
Preparation of Antisera Against Spinach
Hydroxypyruvate Reductase. . . . . . .

Immunodetection of Hydroxypyruvate Reductase .

vii.

84
85
9O
90
90
91
92
93
95
95

99

106

. 111

. 114

118

. 119

. 124

124
125

. 125

. 126

. 128

. 129
. 130

Results 0 C O O O O O O O O O O O O O O O O 0 O O O O O

Hydroxypyruvate and Glyoxylate Reductase Activities

in Green and Blue-green Algae. . . . . . . . .
Partial Purification of the NADflzflydroxypyruvate

Reductase and a NADPflzclyoxylate Reductase from

Chlamwdbmonas. . . . . . . . . . . . . . . . . . .
Molecular weight of the NADH:Hydroxypyruvate Reductase

Substrate Specificity of the Reductases. . . . . .
K values. . . . . . . . . . . . . . . . . . . . .
pH Activity Profiles for the Reductases from
Chlamydhmanas. . . . . . . . . . . . . . . . .
The Effect of Anions on Hydroxypyruvate Reductase
Activity . . . . . . . . . . . . . . . . . . .
Immunodetection of NADH:Hydroxypyruvate Reductase.

Discussion . . . . . . . . . . . . . . . . . . . . . .
ERMNMRY AND DISCUSSION. . . . . . . . . . . . . . . . . . . .

Conclusions. . . . . . . . . . . . . . . . . . . . . .

Future Research. . . . . . . . . . . . . . . . . . . .
APPENDIX: Papers, Abstracts, and.Manuscripts in Preparation. .

LIS'r OF W. O O O O O O O O 0 0 O D O O O O I O O O 0 0

viii

. 131

. 131

136
141
142
143
145

154
154

156

161

162

164

169

. 171

Table

Table

Table

Table

Table

Table

Table

Table

Table

Table

Table

10.

11.

LIST OF TABLg

Types of Fermentation Exhibited by Unicellular
MaryoticAlgae..........

O O O O O O O C O 0 O 5

Organisms Reported to have D—Lactate Dehydrogenase
mtiVityO O O O O O ...... O O O 0 O O 12

The Effect of Killing Techniques on the 14C-Labeled
Productsofazlamdomonasreinbardtii. . . . . . . . . .43

Determination of the Stereospecific Configuration of

the Lactate Produced by Chlamdanonas minhandtii (a)

and Formation of D-Lactate by the Pyruvate Reductase
ofChlamydomonas(b)...................46
Estimtes of the Average Intracellular pH of

Chlamydomonas reinhandtii During Aerobic (Light and
Dark)andAnaerobic (Dark) Conditions . . . . . . . . . . 48

Purification Procedure for the Pyruvate Reductase
fromChlanzydomonasreinbatdtii ............56

Smmary of Some Properties of the Pyruvate Reductase
fmmChlamydomonasreinhazfltii. . . . . . . . . . . . . . 61

Phosphate Activation of the Pyruvate Reductase from
Chlmm reimtii O O O O C O O O O O O O O 0 I O 73

Effects of Various Compounds on the Pyruvate Reductase
from Chlamdomonas reinhardtii. ....... . . 76

A Comparison of Mitochondrial Respiration Between Wild
Type Chlamydanonas reinbardtii and the dk97 Mutant

Deficient in Cytochrome Oxidase . . . . . .101

Effect of SHAM and Aminooxyacetate (AOA) on the Rate
of Glycolate Excretion by Air-grown Chlamydomonas

reinhardtii Wild Type and de7 Cells. . . .103

ix

Page
Table 12. The Percent Distribution of 14{C in Glycolate, 'ICA Cycle
Intermediates , and D—Lactate in Air- Chlamydomonas
dk97 Cells After Photosynthesis with [ CJNal-IOO in

either the Presence or Absence of 5 w SHAM . .3. . . . .112

Table 13. Levels of Hydroxypy'nxvate and Glyoxylate Reductase
Activities in Crude Extracts of Chlamydanonas
reinhardtii and a Comparison to the Rate of 1
Photosynthesis and Other Enzymes of the Oxidative
Photosynthetic Carbon Cycle in These Cells. . . . . . . .132

Table 14. NADflzflydroxypyruvate Reductase Activities in Extracts
from High (5%) CD -Grown or Air-Grown Green and Blue-

Green Algae O O 020 O O O O O O O O O O O O O I O O O 0 0134

Table 15. Purification Procedure for the NADflzl-Iydroxypyruvate
Reductase from Chlamdanonas minbardti'i. . . . . . . . .137

Table 16 . Properties of the Hydroxypyruvate and Glyoxylate
Reductase Activities of (1th minhamltii . . . .144

Table 17. The Effect of pH and Phosphate on Some Kinetic
Parameters of the NADflzHydroxypyruvate Reductase of
ChlamydomonaslSl

Figure

Figure

Figure

Figure

Figure

Figure

Figure

Figure

Figure

Figure

Figure

10.

11.

LIST OF FIGUR_ES_

Page
Pathways of Anaerobic Metabolism in Chlamydomonas
minbaIdtiio 0 O O O O O 9 O O O O O O O O O O O O O O O 10
The Oxidative Photosynthetic Carbon Cycle or C2 Cycle
m klatw mum” I I O O O O O O O O O O I O O O O O 23
Distribution of 14C in Methanol-Soluble, Nonvolatile
Products of ailmdanonas reinhardtii Under Decreasing
02 Cementmtions. O O O O O O O O O O O O O O O O O O O 41
Separation of the Pyruvate Reductase from the NADH-
Dependent Hydroxypyruvate and Glyoxylate Reductase
Activities of Chlamydanonas reinbardtii by Elution
fm 8 SGPhflCI'Y]. 8-300 0011.]!!!- o o o o o o o o o o o o o 58
pH Profile for the Partially Purified Pyruvate
Reductase from Chlamydamonas minhatdtii . . . . . . . . 63
Double Reciprocal Plots for the Pyruvate Reductase from
Chlamydomonasreinhatdtii................66

pH Dependence of Substrate Inhibition by Pyruvate of the
PyruvateReductase................... 69

pH Profile of the Pyruvate Reductase from Crude Extracts
of Chlamydomonas reinbardtii in the Both the Presence
andAbsenceofPhosphate................72

Possible Fates of D—Lactate in Chlamydomonas reinhardtii 87

Turnover of D-[14C]Lactate in Air— and 5% (DZ-grown

Inhibition of D-[14C]Lactate Turnover in SHAM-treated

Chlamydomonas dk97 Cells Deficient in Cytochrome Oxidase
Activity........................108

xi

Figure

Figure

Figure

Figure

Figure

Figure

12.

13.

14.

15.

16.

17.

The Percent Distribution of 14C in Glycolate in
Chlamydamonas Wild Type and dk97 Cells in the Presence
andAbsenceofSHAM..................

tx-Keto Acid Reductase Reactions for Hydroxypyruvate,
Glyoxylate and Pyruvate in Chlamydomonas reinbandtii .

Separation of the NADPflzGlyoxylate Reductase from the
MADE-dependent Hydroxypyruvate , Glyoxylate and
Pyruvate Reductase Activities from Chlamdomonas
reinbardtii by Affinity Chromatography on an Affi-Gel

Elm 001m 0 O O O O O O O I O O I O O O C O O O O O O 0

pH Profile of the NADflzfiydroxypyruvate Reductase from
Chlawdanonasreinbamltn

Substrate Inhibition of the NADH:Hydroxypyruvate
Reductase by Hydroxypyruvate and the Partial Relief
of This Inhibition in the Presence of Phosfliate. . . .

pH Profile of the NADH- and NADPH-Dependent Glyoxylate
Reductase Activities from allamydomonas minbazdtii. .

xii

Page

. 110

. 121

139

147

. 149

O 153

ADP

AMP

AOA

ATP

Bis-Tris

BSA

Caps

Ches

Chl

Ci

CoA

EDTA

Epps

Hepes

LIST OF ABCBREVIATIONS

Adenosine 5’-diphosphate
Adenosine 5’-monophosphate
Aminooxyacetate

Adenosine 5’-triphosphate

[Bis(2-hydroxyethyl)imdno-]tris-
(hydroxymethyl)methane

Bovine serum albumin
Cyclohexylaminopropane sulfonic acid
2-[N-cyclohexylamino]ethanesulfonic acid
Chlorophyll

Curies

Coenzyme A

Counts per minute
2,6-dichlorophenolindophenol
5,5-dimethyl[2-14C]oxazolidine-2,4—dione
5.5’-dithiobis-(2-nitrobenzoic acid)
Dithiothreitol
Ethylenediaminetetraacetic acid

N-(2-hydroxyethyl)piperazine-N’-
3-propanesulfonic acid

N-Z-hydroxyethylpiperazine-N’-2~ethane-
sulfonic acid

xiii

MOPS

NADH

NADPH

SDS
SHAM
TCA

Tris

Immunoglobulin G

Michaelis constant

Lactate dehydrogenase
2-[N—morpholino]ethanesulfonic acid
3-[N-morpholino]propanesulfonic acid
Nicotinamide adenine dinucleotide (oxidized)
Nicotinamide adenine dinucleotide (reduced)
Nicotinamide adenine dinucleotide phosphate (reduced)
Nuclear magnetic resonance
Phosphateebuffered saline

Orthophosphate

pK of the ionizable group on an enzyme
Pounds per square inch

Sodium dodecyl sulfate

Salicylhydroxyamic acid

Tricarboxylic acid cycle
Tris(hydroxymethyl)aminomethane

Maximal velocity of an enzymatic reaction

xiAI

CHAPTER I

LITERATURE REVIEW

A. ANAEEQBIC METABOLISM

Cellular production of lactate is generally associated with anaerObic
glycolysis. For instance, in mammalian muscle cells under conditions of
limiting oxygen, the rate of glycolysis increases to meet cellular energy
demands, and lactate accumulates within the cell (1,2), and lactate
accumulation in ischaemic heart tissue also leads to cellular acidosis
(3). In order for a cell to survive under anaerobic conditions for
extended periods of time, it must be able to control the rate of acid
(lactate) accumulation to prevent cellular acidosis, yet still synthesize
ATP by substrate level phosphorylation at rates sufficient to maintain
essential cellular processes. If high rates of glycolysis are to be
sustained, the cell also requires a large carbohydrate supply.

A variety of survival strategies are employed by cells tolerant of
anaerobic conditions (4,5). Yeast and bacteria can maintain high levels
of glycolysis when supplied with a carbon source in their growth medium,

since they are able to excrete accumulated end products (4). Most animal

1

species tolerant of anaerobic conditions adapt to low 02 stress by
reducing the energy demands of the cell to a level which can be met by
low rates of fermentation (4). Such organisms are capable of decreasing
their metabolic rate and inactivating non-essential cellular processes
during anoxia. Mechanisms by which this can be accomplished.have been
extensively reviewed by Storey (4).

A variety of photosynthetic organisms can also adapt to tolerate long-
term anaerobiosis (6,7). Although photosynthetic organisms are normally
considered to be aerObic organisms, there are a number of instances when
these organisms may encounter conditions of low 02. During flooding, the
air spaces in soil become filled with water, and both plant roots and
soil algae may face 0 shortages (6,7). Several aquatic species of plant

2

and algae grow in stagnant ponds or at water depths where 02 concen-

trations are low. Additionally, rice and swamp and.marsh species often
grow in poorly aerated soils, so that the roots, lower stems and
occasionally the entire plant may be submerged. Many of these species

have developed unique morphological features allowing the transport of 02

from the upper parts of the plants to the submerged portions or have

internal gaseous lacunal systems for storage of O2 and 002 (6-8). There

are also photosynthetic bacteria which are obligate anaerobes that use

H S, H

2 2 and other reduced inorganic compounds as a source of reducing

equivalents (9).

When the roots of flood-tolerant plants are initially confronted with
hypoxic conditions, Lelactate is formed as the end product of glycolysis
(6,10,11). Hewever, after a short period, the cells have the capability
of switching their metabolism from lactate production to ethanol
fermentation, avoiding excessive acid accumulation in the cell
(6,10,11). USing both 31P— and 13C-nmr with intact maize root tissues,
Roberts et a1. (11) have demonstrated that during the period of initial
lactate formation, the cellular pH decreases to around pH 6.5. As the pH
decreases, ethanol begins to accumulate in the root tissue cells. From
the results of in vitro kinetic data, Davies (10,12) proposed that this
decrease in pH approaches the pH optimum.of pyruvate decarboxylase, the
enzyme catalyzing the first step of ethanol formation from pyruvate. The
decreasing pH moves away from the pH optimun of the L-lactate
dehydrogenase and, at lower pH values, ATP inhibition of lactate
dehydrogenase is more severe (12). Thus, conditions favor the formation
of acetaldehyde from.pyruvate, via pyruvate decarboxylase, over lactate
formation. The acetaldehyde, in turn, is converted to ethanol in a
reaction catalyzed by alcohol dehydrogenase (6,10,11). Mutants deficient
in ethanol formation (i.e. alcohol dehydrogenase mutants) are unable to
tolerate extended periods of anaerobiosis. Nmr studies with root tissues

from such mutants indicate that uncontrolled lactate production occurs in

these cells, resulting in cellular acidosis (11).

(1"

Flood-tolerant species have additional features allowing them to
survive during extended periods of anaerobic conditions. Ethanol, which
can be toxic to the plant, can either be excreted from plant roots or
transported up to aerobic portions of the plant where it may be oxidized
(6). Alternatively, some species diversify their glycolytic pathway to
form end products which are less toxic, such as malate and alanine
(6,13). Plants also have large carbohydrate reserves, enabling them.to
sustain rates of fermentation sufficient to meet the energy demands of
the cell for long time periods.

Many unicellular algae are also tolerant of anaerobic conditions.
various anaerdbic fermentation pathways have been shown to exist in algae
(14,15), which can be classified.into three main types (Table 1). Algae
from the first classification form lactate exclusively during starch
fermentation, resembling the homo-lactic fermenting bacteria (14-16).

The second class of algae produce ethanol with little or no anaerobic
accumulation of acids, as do both yeast and some plant roots (6,14,15).
The third classification is hetero- or mixed-acid fermentation. Algae in
this group form a variety of anaerobic products, including acetate,
formats, ethanol, lactate, and H2, and occasionally glycerol and
butanediol (14,15,17—19).

Algae from any of the three classes may also evolve CO2 and H2 during
anaerobic metabolism (14,15,17-22). Many unicellular algae have the

ability to utilize and/or evolve molecular hydrogen due to the induction

Table 1. Types of Fernentation Exhibited by Unicellular Buharyotic Algae.

Anaerobic Starch Representative References
Detradation Products Species
I. Bonn-lactic Lactate, COO, Hz Chlorella pyrenoidosa 14-16
Feraentation “ Scenedesnos obliquue

Anhistrodeeaus braunii
Prototheoa zopfi
Cyanidina caldariua

11. Alcohol Ethanol, little or Chlorella ellipsoidea 14,15
Perlentation no lactate, CO , H Chlorella sp. Harburx
2 2 . .
Cblorella nrnrata
Ochroaonaa aalhaaensis

III. Mixed-Acid Acetate, forlate. Chlaaydononae reinhardtii 14,15,
(hetero-l ethanol, glycerol, Cblaaydoaonas noevueii 17-19
Fernentation lactate, CO , H , Cblorella vulgaris

butanediol Chlorella fusca

Cblorozoniua elongatua
Coelaetrua proboscideun

of the synthesis of hydrogenase under anaerobic conditions (20-21). This
system provides energy for the cell, and under certain conditions, even
allows for anaerdbic 002 fixation.

Krequerg (18) reported that 97% of the fermentation products of
Chlamydanonas reinhardtii, Chlorogonium elongatum, and Chlorella fusca
were excreted into the medium. Since the algae do not accumulate these
end products intracellularly, and the cells have large carbohydrate
reserves (starch), they are able to tolerate anaerobic conditions.
Hewever, only low rates of anaerobic starch fermentation (approximately
100 to 300 thmes less than the rates observed for bacteria and yeast)
have been observed in algae (15). Although such low rates of
fermentation apparently provide sufficient energy for maintaining
essential cellular functions, not enough ATP is synthesized to sustain
cell growth during anaerobiosis (15).

Anaerobic lactate production has been reported for a number of
unicellular algae. In only one algae, Chlorella fuses, was the L-isomer
reportedly formed (17). warburg (23) first reported the anaerdbic
accumulation of D-lactate in Chlorella. Since then, D-lactate has been
observed in the green algae Chlamwdbmonas reinhardtii (24), Chlamydbmonas
moewusii (17), Chlamydomonas sp. IAMrC-Zl (25), Sbenedesmus obliquus
(26), Sbenedesmus basilensis (25), Chlorella vulgaris (14,25), and

Chlorella pyramoidbsa (14), in the colorless alga, Prototheca zopfi (14),

and in the blue-green eukaryotic alga, Cyanidium caldarium (16).

 

B. ANAEROBIC METABOLISM IN CHLAMYDOMOM4§

Chlamydbmonas is a primitive unicellular eukaryotic green alga of the
order Volvocales, class Chlorophyceae (27). This algae is commonly found
in soil, both fresh and marine water, and even in anaerobic waste-
stabilization ponds (27,28). The flagellated cells are bounded.by a cell
wall and possess a single, large cup—shaped chloroplast (27). The
genetics of ChlamydbmonaS'are well characterized, making it a useful
organism for metabolic studies and for the production of mutants.

Since Chlamydbmonas often grows in soil, it is probable that the
cells would encounter anaerobic conditions during the flooding of soil
air spaces (7). Indeed, these algae seem well adapted to survive during
extended periods of anoxia. The cells have large starch reserves to
sustain fermentation and have an inducible hydrogenase system (17—
19,21,29,30). In fermentation studies, after 3 to 6 hours of
anaerobiosis, the starch degradation products of Chlamydomonas
reinbardtii are formats, acetate and ethanol in a 2:1:1 ratio, and
smaller amounts of 002 and H2 gas (18,19). ChlamydOmonas moewusii also
exhibits mixed-acid fermentation, but the major product from starch
degradation is glycerol (17). C3 moewusii also produce significant

amounts of 002, acetate, H2, ethanol, D—lactate, butanediol, and formats.

The pathways of anaerobic metabolism,in.Chlamydomonas reinhandtii are
summarized in Figure 1. The regulatory enzyme of the mixedracid
fermentation pathway is apparently pyruvate formats-lyase which catalyzes
the formation of one molecule of acetyl-CoA and one molecule of formats
from pyruvate and coenzyme A (18,31). Although present constitutively,
the enzyme exists in a catalytically inactive form in aerobic cells.

When cells become anoxic, the enzyme is activated by a complex mechanism
requiring reduced flavodoxin, S-adenosybnethionine, pyruvate (as an
allosteric effector), and an Fe-dependent activase (31). This enzyme was
previously thought to occur only in prokaryotic cells.

The acetyl-CoA produced in the pyruvate formats-lyase reaction has at
least two possible fates (18,19,31). Aldehyde dehydrogenase catalyzes
the reduction of acetyl-CoA to acetaldehyde, which in turn is reduced to
ethanol by alcohol dehydrogenase (Figure 1). Secondly, acetyl-CoA can be
converted to acetate in an ATP-producing pathway (Figure 1).
Phosphotransacetylase catalyzes the formation of acetyl-phosphate, a high
energy compound; subsequently, acetate and ATP are formed in a reaction
catalyzed by acetate kinase. In the presence of hypophosphite, an
inhibitor of pyruvate formats-lyase, or after excessively long periods (8
to 28 hr) of anoxic conditions, Chlamydomonas apparently produce ethanol

via pyruvate decarboxylase and alcohol dehydrogenase (18).

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Although not observed as a long-term starch fermentation product in
Chlamydbmonas reinhandtii, anaerobic lactate production has been reported
for these cells (17-19,24). Additionally, when hypophosphite was added
during the long-term fermentation studies to inhibit the pyruvate formats-

lyase, Delactate, OO and ethanol were produced rather than formats and

2
acetate (18) . The production of D-lactate by Cblamydomonas reinhardtii

during short time periods of anaerobiosis is described in Chapter II.

C. D-LACTATE DEHYDROGENASES

unicellular Green Algae. The presence of lactate dehydrogenases
specific for the D-isomer of lactate has been reported for a variety of
unicellular algae (24,32—35). Delactate reportedly can also be oxidized
by a mitochondrial membrane-bound.dehydrogenase, commonly referred to as
a glycolate dehydrogenase (36-42). This enzyme catalyzes the oxidation
of glycolate to glyoxylate in the oxidative photosynthetic carbon (CZ)
cycle (see below). Algal lactate dehydrogenase activities are discussed
in depth in Chapters III and IV. The unicellular algae with reported
D—lactate dehydrogenase activity are listed in Table 2.

Yeast and Fungi. The D-lactate dehydrogenase of yeast, a zinc-
requiring flavoprotein, is linked to electron transport (48). In vitro,
the dehydrogenase will transfer electrons only to cytochrome c or to

phenazine methosulfate. This enzyme is distinct from both a L-lactate

dehydrogenase (cytochrome b?) and a D—u-hydroxy acid dehyrogenase of

 

Table 2. Organiser Reported to

1.2

have D-Lactate Dehydrogenase Activity.

References are given in

 

parenthesis.
A. The linxdoa Protista
1. Phylcl Chloropbycopbyta 2. Phylcl Rutlenopbyta (35,39,10)
Rutlena (rsciiis
a. Class Chlorophyceae‘
1) Order Volvocales
Chlasydoaonas reinhardtii (24,36) 3. Pbylnn betridiolycota (43-45)
Chlanydosonas sp. Ilfl-C-Zl (33) Chytridiun
Dunaliella tertiolecta (32,37) Hacrochytriua
Blastocladia
2) Order Chlorellales Blastocladiella eaersonii
Chlorella vulxaris (23.32.37) Allosyces arbuscula
Cblorella pyrenoidosa (32,37)
Bresosphaera viridis (32,34,37,38)
Oocytis polylorpha (32,37) 4. Phylul Phycoaycota (43-45)
a. Class Ooaycotea
3) Order Chlorococcales (37,41) Saprosyces (2 species)
Chlorococcua (5 species) Bhipidius
Neochloris aquatics Hindeniella spinospora
Neospongiococcua (5 species) Aqualinderella feraentans
Protosiphon botryoides Saprolegnia (2 species)
Achlya (2 species)
4) Order Chlorosarcinales (41) Pythiua debaryanul

Axilosphaera vegetata
Borodinellopsis terensis
Chlorosarcina (2 species)
Chlorossrcinopsis eresi
Planopbila terrestris
Tetracystic (2 species)

3 . .
Classizied as described by Sold and iynne (2?).

Aphanosyces euteiches
Thracstotheca clavata
Apodacblya (2 species)

b. Class Hyphochytridionycotea
Rhizidionyces apopbysatus

1.3

Table 2 (continued)

5) Order Caulerpales (37) 5. Phylun Sarcodina (46)
Codius sp. a. Class Acrasea (slice colds)
Polyspondyiiua pallidua
6) Order Chaetophorales (37)
Stizeocloniua helveticua

7) Order Ulotrichales (32,37)
Hicrospora sp.

b. Class Prasinophyceaeb (42)
Asteroaonas sp.
Heterosastir sp.
Hicrononas (2 species)
Pedinosonas sp.

Platynonas sp.
Pyrasisonas (3 species)

b
Bold and Wynne (27) do not consider Prasinophyceae as a separate class in the phylua
Chlorophycopbyta.

2141

Table 2 (continued)

B. The lingdon Fungi D. The Kingdoa lnisalia (50,51)
1. Phylan Deuteronycotina 1. Pbylun lrthropoda
('Funsi Ilperfecti') a. Subphylua Chelicerata
a. Class hyphonycetes ((7) 1. Class Herostoaata
Piricularia orysae hisulus polyphesus

(horseshoe crab)
2. Pbylua hscosycotina
a. Class Teaiascolycetes ((8) 2. Class Arachnida
(yeasts) Pholcus phalantioides (spider B)
Dugesiella hentsi (tarantula)
Geolycosa (wolf spider)
Centruroides sculpturatus
(scorpion)
C. The lingdos lonera Tetranychus pacificus (bran site)
Opiliones (daddy lont less)
- The Buhacteria Category ((9)
b. Subphylul Handibulata

Lactohacillus (18 species) 1. Class Crustacea

Leuconostoc (6 species) Balanus nubilus (acorn barnacle)
Streptococcus (2 species) Tetraclita squasosa rubescens
Pediococcus (4 species) (cone barnacle)

Aerobacter (2 species) Pollicipes polyserus
Escherichia (2 species) (xoosenech barnacle)

Hafnia sp. Penella (fish copepod)
llebsiella sp.

Salnonella typhisurius 2. Phylua Hollusca

Proteus vulgaris a. Class Gastropoda

Serratia sp. Polinicies heros (noon snail)
Acetohacter perorydans Helix aspersa (garden snail)
Pseudosonas (2 species) Haliotus (3 species - abalone)
Propionibacteriua pentosaceus Crepidula (slipper shell)
Staphylococcus (nuaerous species) Acnaca pelts (shield liapet)
Hyeoplasna (nunerous species) Hegathura crenulata (giant
Butyribacterius rettgeri keyhole linpet)

Selenoaonas rusinantiul Aplysia californica (see here)

Table 2 (continued)

1.5

b. Class Anphineura
Cyanoplar hartsexii (chiton 1)
Nuttalina flura (chiton 2)
c. Class Pelecypoda
Pseudochasa erogyra (rock clan)
d. Class Cephalopoda
Octopus bisaculatus

3. Phylus Annelida
a. Class Polycbaeta
Hereis sirens (Iarine clan worn)

16

anaerobic yeast (48). Although all three of these dehydrogenases are
flavoproteins, their specificity for electron acceptors varies greatly.

Only the lower classes of fungi ferment glucose to lactate; the
higher fungal species convert glucose to ethanol and 002 (43). Pyridine
nucleotide-dependent lactate dehydrogenases, specific for the formation
of D—lactate, have been reported for a number of these species (Table 2).
It should be noted that the lower fungi, including the phyla
Chytridiomycota and Phycomycota hays been reclassified under the Kingdom
Protista and are no longer considered true fungi (52). The Oomycetes are
even considered by some to be heterotrophic algae, rather than fungal-
like organisms (52). The lactate dehydrogenases from these species are
apparently not associated with electron transport, as is the case with
yeast. Interestingly, only sapromyces, can reutilize the D-lactate they
produce by oxidative metabolism. Ffiricularia, classified.as a "Fungi
Imperfecti" (52), reportedly has both D— and Lelactate dehydrogenases
(47). The slime mold, Polyspondylium pallidium, has also been reported
to have a D—lactate dehydrogenase (46).

The D—lactate dehydrogenase from Pythium debaryanum is allosterically
inhibited by GTP, and GTP inhibition of D—lactate dehydrogenases from a
number of other species in the Oomycete and Hypochytridiomycete classes

had also been observed (45).

17

Bacteria. Both soluble and membrane-bound forms of D—lactate
dehydrogenases have been reported for a variety of bacteria (Table 2,
ref. 49, 53-59). Some bacteria reportedly have both D- and L-specific
lactate dehydrogenases (49,53). The topic of bacterial lactate
dehydrogenases has been reviewed at length (49,59). Many bacteria
produce D—lactate as an end product of anaerobic glycolysis; other
bacteria are able to use D-lactate as a source of both carbon and energy
for growth (49,59). Bacterial D-lactate metabolism has been best
characterized for ES 0011, where D-lactate is a respiratory substrate and
provides energy to drive the uptake of certain growth substrates (59-61).

When Eh coli is grown on glycolate as a carbon source, glycolate
dehydrogenase activity, which is apparently linked to electron transport,
is induced (62). D—lactate oxidation was also observed, but it was
unclear whether or not this reaction was catalyzed.by the same
dehydrogenase as was glycolate. This bacterial glycolate dehydrogenase
had several characteristics in common with the algal glycolate
(D—lactate) dehydrogenase. Previously, Kornberg and Gotto (63)
demonstrated that Pasademonas, growing on glycolate as a carbon source,
could convert glycolate to glyoxylate, indicating that these bacteria
also had.a glycolate-oxidizing enzyme.

Animals. D-lactate dehydrogenases have been reported for several
invertebrates (50,51) and a list of these species is presented in

'Table 2. Mitochondria of mammalian cells possess a zinc—requiring

18

Des-hydroxy acid dehydrogenase which will catalyze the oxidation of
D-lactate (64). The best characterized.invertebrate D-lactate
dehydrogenase is that from the horseshoe crab, Limulus‘polyphemus
(50,65). Both heart and muscle isozymes of the enzyme are present in
L. polyphemus. Tissue specific isozymes have also been observed in the
scorpion and one spider (51). Like most of the D-lactate dehydrogenases,
the isozymes from L. polyphemus are dimers (65,66). This is in contrast
to the L-specific lactate dehydrogenases which are tetramers (66).
Evolutionary Significance. Tolbert (32,37,67) has emphasized the
evolutionary differences between the unicellular algae and the
multicellular algae and higher plants with respect to lactate and
glycolate metabolism. unicellular algae have a mitochondrial membrane-
bound glycolate and/or D-lactate dehydrogenase (36,38-40,68). This
cyanide-sensitive dehydrogenase is apparently linked to electron
transport (Chapter IV, and ref. 39,40,69). unicellular algae produce
D-lactate anaerobically via a soluble D—lactate dehydrogenase or pyruvate
reductase (Chapter III). On the other hand, both multicellular algae and
higher plants have a cyanide-insensitive glycolate oxidase which
catalyzes the oxidation of both glycolate and the L-isomsr of lactate,
transferring electrons directly to molecular oxygen to producing H202
(70). Glycolate oxidase is a peroxisomal enzyme, as is catalase which
degrades the H 0 produced during glycolate oxidation (70). O is not

2 2 2

the direct electron acceptor for the algal glycolate dehydrogenase, so

l9

H202 is not produced during glycolate metabolism in these cells. unlike
unicellular green algae, higher plants have soluble lactate
dehydrogenases which specifically form.L~lactate and are similar to
mammalian Lelactate dehydrogenases (12,71,72).

Long and Kaplan (73) have proposed that both 0- and L-lactate
dehydrogenases evolved.from a single gene and.hypothesize that minor
amino acid substitutions in the active site could alter the spatial
orientation of pyruvate sufficiently to produce a different.
stereospecific reaction product. D- and L-lactate dehydrogenases have
different specifity for pyridine nucleotide analogs (53,74), and
transition state analogs (75,76). D-lactate dehydrogenases do not bind
to oxamate—diaminohexyl-Sepharose columns; whereas this affinity column
is widely used.in the purification of L-lactate dehydrogenases which do
bind this resin (77). Comparison of the active site sequence of the
D-lactate dehydrogenase from Limulus polyphemus with sequences from
L-lactate dehydrogenases revealed no obvious sequence homology (65).
This suggests that D— and L-lactate specific lactate dehydrogenases
either evolved from different proteins or, if D-lactate dehydrogenase

evolved from the same gene as did L—lactate dehydrogenases, there were

major rearrangements in the active site sequence.

20

D. THE OXIDATIVE PHOIOSYN'I‘HEI‘IC CABBON (CZ) CYCLE

This laboratory has been specifically interested in the metabolism
of three intermediates of the oxidative photosynthetic carbon cycle -
glycolate , glyoxylate and hydroxypyruvate . We were also interested in
the possible relationship between D-lactate metabolism and the C2 cycle
for a variety of reasons. D—lactate is a structural analog of glycolate
and reportedly, is also a substrate for glycolate dehydrogenase. This
enzyme was studied with respect to its role in D-lactate metabolism in
Chlamydanonas (Chapter IV). The possibility was examined that the
glycolate (D—lactate) dehydrogenase, and therefore, D—lactate oxidation,
was linked to mitochondrial electron transport (Chapter IV). Unicellular
algae can excrete glycolate into their surrounding medium (70,78), so it
was of interest to see if Chlamydomonas was also capable of excreting
lactate. Hydroxypyruvate and glyoxylate, as well as pyruvate, are
substrates for lactate dehydrogenases from both plant (12,72,79) and
manmalian. (66,80) sources, so it was of interest to see if the pyruvate
reductase (D—lactate dehydrogenase) of Chlamydanonas also catalyzed the
reduction of these two C2 cycle intermediates (Chapter III). It was also
of interest to determine whether Chlamydomonas had other enzymes which
catalyzed the reduction of hydroxypyruvate and glyoxylate, as do higher
plants. Hence, the partial purification and characterization of two

additional enzymes from Chlamydomonas, NADlizhydroxypyruvats reductass and

NADPiizglyoxyrlate reductase, are described in Chapter V.

21

The author and other members of this laboratory have recently
extensively reviewed the oxidative photosynthetic carbon cycle (70), so
only a relatively brief review of this pathway is presented here. The

light-dependent O uptake and 00 release that occurs in photosynthetic

2 2

organisms has been termed photorespiration. The metabolic pathway for

photorespiration, in which sugars are oxidized to CO in the light, is

2

known as the oxidative photosynthetic carbon cycle or C

2

cycle. The C2

cycle coexists with the reductive photosynthetic carbon cycle (C3 cycle)

in which 002 is reduced to sugar phosphates in the chloroplasts and

subsequently converted to starch for storage or transported to the

cytoplasm for energy or sucrose synthesis. The C2 and.C3 cycles are

linked.due to the inevitable competition between 002 ando2 at the

catalytic site of ribulose 1,5-bisphosphate carboxylase/oxygenase, the

enzyme which catalyzes the first step in both 002 fixation and the C2

cycle.
A scheme of the oxidative photosynthetic carbon cycle and closely
related metabolic steps in higher plants is shown in Figure 2. The first

step of the C cycle is the formation of 2-phosphoglycolate from ribulose-

2

1,5 bisphosphate and O by the oxygenass reaction of ribulose-P

2 2

carboxylase/oxygenase in the chloroplasts. This enzyme also catalyzes
the first step of photosynthetic 002 fixation, in which two molecules of
3-phosphog1ycerate are formed from one molecule each of ribulose—1,5—

bisphosphate and 002. A specific phosphatase, also located in the

22

Figure 2 . The Oxidative Photosynthetic Carbon Cycle or C2 Cycle and
Related Pathways. The numbers shown refer to the name of the enzyme or
reaction: (1) ribulose-P2 .carboxylase/oxygenase, (2) P-glycolate
phosphatase, (3) glycolate oxidase, (4) catalase, (5) glutamate/
alaninezglyoxylate aminotransferase, (6) glycine decarboxylase complex,
(7) serine hydroxymethyltransferase, (8) serinezglyoxylate
aminotransferase, (9) NADH:hydroxypyruvate reductase (D—glycerate
dehydrogenase) , (10) glycerate kinase, (11) P-glycerate phosphatase,
(12) glutamine synthetase, (13) glutamate synthase, (14) NADzmalate
dehydrogenase, ( 15) aspartate:o<-ketoglutarate aminotransferase,

(16) NADI-Hzglyoxylate transferase, (17) carbonic anhydrase, (18) photo-
phosphorylation. Translocators of intermediates of the pathways are
indicated by letters: (A) phosphate translocator, (B) glycolate/
D—glycerate translocator, (C) chloroplast dicarboxylate translocator(s) ,
(D) mitochondrial amino acid translocator(s), (E) mitochondrial

dicarboxylate trans locators .

23

 

 

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24

chloroplasts, hydrolyzes P—glycolate, and the resulting glycolate is
transported out of the chloroplasts to be further metabolized. In leaf
peroxisomes, glycolate is oxidized by the flavoprotein glycolate oxidase

producing glyoxylate and H Catalass degrades the H O formed.during

202° 2 2
glycolate oxidation, and the glyoxylate is transaminated to glycine. In

the mitochondria, one glycine molecule is oxidized to 002, NH3, and a

Cl-unit bound to tetrahydrofolate. This Cl-unit is, in turn, added to a
second glycine molecule to form serine. A.peroxisomal aminotransferase
converts the serine to hydroxypyruvate, which in turn, is reduced to
D-glycerate. The glycerate can return to the chloroplast, where its
phosphorylation is catalyzed by glycerate kinase. The resulting
3-Pbglycerate can reenter the reductive photosynthetic carbon cycle to
regenerate ribulose-P2 or, after reduction to triose—P in the
chloroplast, can be used for synthesis of storage carbohydrates.

Since the C2 cycle involves the synthesis and metabolism of
glycolate, the term.glycolate pathway has been used for the irreversible
portion of the cycle involving glycolate biosynthesis and subsequent
conversion to glycine. The term glycerate pathway refers to the
reversible part of the pathway between serine and P-glycerate (67).

Other important reactions closely associated with the C2 cycle include
the shuttle of reducing equivalents between subcellular compartments, and
the photorespiratory nitrogen cycle which functions to reassimilats NH3

released by photorespiratory glycine oxidation.

25

During photorespiration in higher plants, 0 uptake occurs during the

2

formation of P-glycolate via the oxygenass reaction of ribulose-P2

carboxylase/oxygenase (reaction 1) and during glycolate oxidation in the
peroxisomes (reaction 3). Also, NADH produced in glycine oxidation might
be utilized.by the mitochondrial electron transport chain, resulting in

glycine-dependent O uptake» The primary site of 00 release in the C

2 2 2

cycle occurs during glycine oxidation in the mitochondria (reaction 6).
The functions of the oxidative photosynthetic carbon cycle

(photorespiration) have long been debated (67,81—83). The pathway is

considered to be a wasteful energybburning process which occurs because

of the oxygenass reaction associated with ribulose-P2

carboxylase/oxygenase. However, because of the ability of the C2 cycle

to dispose of excess photosynthetic assimilatory capacity, it has been
proposed that the cycle prevents photooxidative damage, especially during

periods of stress-induced CO shortages (84,85). Since the oxygenass

2

activity that occurs in standard atmospheric conditions may be an

inevitable consequence of the ribulose-P2 carboxylase/oxygenase catalytic

mechanism (81), a mechanism for the metabolism of the product P-glycolate

must exist in the plant. This is accomplished by the C cycle at the

2
expense of losing at least 25% of the carbon as CO2 (83). However, three

of four carbon atoms flowing through the C2 cycle reenter the C3 cycle

and some of the released 002 can be photosynthetically refixed by the

cell. Hence, the C2 cycle may be viewed as a carbon scavenging system.

26
An additional function of the C2 cycle may be to provide glycine and

serine, and Cl-units which are required for numerous biosynthetic

pathways (67,86).

Although well characterized in higher plants, much less is known
about the C2 cycle in unicellular algae. However, some differences in
the algal C2 cycle are already known. As noted.abcve, the enzyme
catalyzing the oxidation of glycolate to glyoxylate in algae is a
mitochondrial membrane-bound dehydrogenase, rather than a peroxisomal
oxidase as in higher plants (70). Other enzymes of the C2 cycle which
are located in the peroxisomes of higher plants, including the

aminotransferases and hydroxypyruvate reductase, are mitochondrially

located in the green algae (70).

CHAPTE! II. D—IACI‘ATE NATION IN W WI

27

28

INTRODUCTION

The anaerobic production of D—lactats by unicellular green algae was
first noted in Chlorella by Warburg (23). D-lactate has also been
observed in various other algae (14,16,17,24—26). Tolbert et a1. (87)
showed.that if Chlamwdcmonas*were pelleted by rapid.centrifugation after
2 minutes of photosynthesis, 51% of the newly fixed 1400 was found in

2
lactate in cells which had.been grown with air enriched.with 5% CO and

2
26% in cells which had.been grown with air levels of 002 (0.04%).
Despite this evidence for the formation of D—lactate, the metabolic role
of this compound in algae is not known.

Studies of anaerobic metabolism in algae have concentrated on
quantitative analyses of starch fermentation products formed after
several hours of anaerobiosis. The results indicate that various
fermentation pathways exist in algae (Table 1); Chlamydomonas reinhardtii
exhibits mixedeacid fermentation. The long-term (3 to 6 hr) anaerobic
starch fermentation products of Chlamydomonas reinhardtii include
formats, acetate, ethanol, and smaller amounts of 002 and H2 (18,19).

D—lactate was observed in these long-term studies only at extreme pH

values or when the pathway of formats formation was inhibited by

29
hypophosphite (18) . However, little is known as to what metabolic
changes occur during the initial minutes of anaerobic conditions. The
results described in this chapter demonstrate the rapid accumulation of
D-lactate in Chlamydomonas reinbardtii during short-term anaerobic

experiments of a few seconds to 30 minutes.

30

MATERIALS AND PERIODS

Materials

dalmdomonas reinhardtii, UTEX 90, Anabaema variablis UTEX B 337,
and Synechococcus leopoliensis UTEX 625 were from the R. C. Starr
collection of the University of Texas, Austin. The F-60 mutant of
Chlamydanonas reinhandtii was a gift from Dr. R. K. Togasaki, Indiana
University.

[Momma was from New England Nuclear. The silicon oils AR 20 and
AR 200 were provided by Wacker Chemie, thich, F.R.G. Cannon laboratory

chemicals were of reagent grade, and solutions were prepared in

deionized, distilled water .

Growth a_.nd Harvesting of Algae

Wildtype Chlamydomonas were grown at 23-25°C in a high phosphate
medium with (NH4)NO3 at pH 6.8 (88). The F-60 mutant was grown at pH 7.3
in Tris-acetate-phosphate media (89) . The blue-green algae were grown at
pH 8.5 in the media of Kratz and Myers (90) with the addition of 10 um
sodium N-tris[hydroxymethyl]-2—aminoethanesulfonic acid. The cells were
aerated with either air (0.04% 002) or with air enriched with 5% CO2 in

flat Erbach flasks and were continuously illuninated under 125

-2 -1
)lEinsteins-m see from cool white fluorescent tubes.

31

Cells were harvested after 3 to 4 days, near the end of the log phase
of growth, by centrifugation at 900g and 4°C for 5 min, washed once with
distilled water, and recentrifuged in tared Corsx tubes at 12,000g for 10
min. The algal wet weight was recorded and the pelleted cells were
resuspended for photosynthetic experiments in either 3 mM potassium
phosphate buffer, pH 7.5, or 25 m Hepes buffer at pH 7.5. The algal
suspensions were stored on ice, and all photosynthetic experiments were
done within 1 hr after harvest. The chlorophyll concentration of algal
cell suspensions was measured spectrophotometrically after acetone
extraction (91).
14C-I..abeli of as

An aliquot of resuspended Chlamydomonas cells (either 2 or 5% w/v)
was placed in a 2 cm diameter glass vial which was held in a plastic
holder in a circulating water bath at 25°C. The contents were stirred by
a magnetic bar. The samples were preilluninated for 2 min by 1000
)lEinsteinsom-z-sec“1 of light from a projector. After this light

adaptation, [14C1NaHCO was added to the suspension to a final

3

concentration of 1 11M. When illumination times were longer than 5 min,

additional [MCJNaI-IOO:3 was added to 1 m at 5 min intervals to ensure a

saturating level of inorganic carbon for the cells. Aliquots of the cell

suspension were removed at various time points and CO2 fixation was

stopped by one of two methods. In one procedure designed to assay the

u

on
‘1

32

cellular components and the surrounding medium separately, the cells were
imediately separated from the suspension by centrifugation for 5 s in an
Eppendorf microfuge. The supernatant was removed with a Pasteur pipette
and added to an equal volume of methanol. The pellet was resuspended in
50% methanol to a volune equal to the original volume of the aliquot in
order to terminate enzymtic reactions and to solubilize the cellular
contents. In the other procedure, an aliquot of the photosynthesizing
cells was rancved and directly added to methanol and mixed, without cell
separation from the medium.

To count radioactivity in these samples, 50 )11 were added to 0.45 ml
of 0.5 N acetic acid and allowed to sit for 3 hr to release unfixed 14C.

Scintillation fluid (4.5 ml) was then added, and the radioactivity was

determined in a liquid scintillation spectrometer.

Two-Dimensional Paper Chromatoggggyz

Aliquots of the cell and supernatant samples from 14C-labeling
experiments were chromtographed for product identification as previously
described (92) . The radioactive compounds were located by exposure of
the chromatographs to Kodak XAR-S X-ray film for about 3 weeks . To
determine the percentage of label in each compound, the areas
corresponding to spots on the X-ray film were cut out of the
chromatograms and into approximately 5 m2 pieces and placed into

scintillation vials. Compounds were eluted from the paper by adding

33

1.5 ml of water to each vial, followed by gentle mixing for about 2 hr.
Scintillation fluid was then added, and the samples were mixed and the
radioactivity counted.

Lactate was identified by cochromatography with a [14C]1actate
standard and with an unlabeled lactate standard. The unlabeled carrier
was located on the chromatograms by spraying them with a 0.04% solution
of bromocresol green in ethanol adjusted to pH 7.0. Lactate was also
identified by gas chromatography-mass spectrometry of the trimethylsilyl
derivatives of the organic acid fraction from the algal cells using the
procedure described by DeWitt et a1. (93) . In this case, identification
was based upon both the retention index and sass spectrun. Attempts to
identify lactate by the colorimetric test reported by Warburg (23) , using
2,7-dioxynapthalene, did not give a red color as previously indicated;
rather, a yellow-green solution was observed, and the absorbsnce at 460

nm was not proportional to the concentration of lactate in the samples.

AnalEis of the Products of MODE-Fixation by Cthomonas in Decreasing

92 Concentrations

A 5% (w/v) suspension of high—CO grown cells (approximately 100 to

2
120 )1g Chl/ml) was placed in a Rank Brothers (Bottingham-Shire) O2
electrode chamber for continuous monitoring of the 02 concentration. The

cells were illuminated with 800 )-IEinsteins-m-2-secm1 of light from a

1
projector and [ 4CJNaHC03 (0.25 mCi/mnole) was added to a final

34

concentration of 1 mM. The cells were allowed to photosynthesize for 2
to 4 min, and then the light was turned off and the chamber covered with
aluminum foil. In the dark, the system became anaerobic within 5 to 10
mun.depending on how high the 02 concentration reached.during
photosynthesis. Throughout the experiment, 200 #1 samples were removed
and iumediately added to 500 )ll of methanol to terminate reactions.
After each aliquot was removed, the chamber cap was adjusted to prevent

formation of air bubbles in the system. The samples were concentrated by

evaporation to around 0.1 m1 and analyzed by paper chromatography.

Deteggigation of the Stereogpggific Configuration of the Lactate Produced

Chl omonas reinhandtii

A suspension of 5% OOZ-grown ChlamydomonaS'in phosphate buffer, pH
7.5, were allowed to photosynthesize with Nah-112003 which was added to 1
mM at 0 and 15 min. After 30 min, the cells were pelleted by
centrifugation at 7700g for 5 min at room temperature to produce
anaerobic conditions. Reactions were terminated by resuspending the
pelleted cells in 5.8 N perchloric acid. The sample was neutralized with
KOH.and.centrifuged. Bacterial NAD:D~lactate dehydrogenase
(Lactobacillus lsiohmannii) and porcine heart L-lactate dehydrogenase
'were used to determine the presence of either D— or L-lactate in the

resulting supernatant sample according to the method of Gawehn and

Bergmeyer (94). The reaction mixture (pH 9.0) contained 453 mM glycine,

35

352 mM hydrazine hydrochloride, and 2.47 mM NAD. An aliquot of the
supernatant (1.1 ml) was added to 3.3 m1 of assay buffer and 0.5 m1 of
this was used.for each assay. After mixing, the initial absorbence at
340 nm was measured. The reaction was initiated with the addition of
either D-LDH or L-LDH to a concentration of 60 units/ml (units were
Pmoles of NAD reduced/min at 25°C). After 90 man at 25°C, the absorbsnce
at 340 nm was compared with a control which contained no algal sample.

No further increase in absorbence was detected after this 90 min time

period.

Formation of D—Lactate by the Ezguzate Reductase of Chlamydomonas

An enzyme fraction was prepared.from 5% OO -grown Chlamvdomonas'by

2
extraction in 50 mM potassium phosphate, pH 7.0, containing 3% Triton
X-100, 1 mid 171T, and 5 m EDTA for 40 min, followed by centrifugation.
The supernatant was used to prepare a 45 to 80% saturated (NH4)ZSO4
fraction. The protein was resuspended in 3 mM potassium phosphate
buffer, pH 6.2 and used for pyruvate rsductase assays. No enzyme
fraction was added to the "blank", and the "control" assay was run
without pyruvate. The assay contained potassium phosphate buffer, pH
7.0, 0.12 mM NADH, and 30 P1 enzyme fraction. The reaction was initiated
with 2 mM pyruvate and incubated for 90 min at 25°C, and then was

terminated by protein precipitation with 5.8 N perchloric acid. After

neutralization with KCH and centrifugation to pellet the protein and

36
perchloric acid, the supernatant samples were used as substrates in

assays with the stereospecific lactate dehydrogenases as described above
to determine the configuration of the lactate produced in the pyruvate

reductase reaction.

Estimates of Intracellular Hg

The intracellular pH of 5% CO -grown Chlamydomanas reinhardtii was

2
determined by measuring the partitioning of the weak acids [14C1benzoic
acid and [140mm between the cells and the incubation medium as
described by Gehl and Colman (95). After incubation with the 14C—labeled
weak acid, 200 Pl aliquots of the samples were removed.and layered as the
top layer of prepared silicon oil tubes. These tubes were prepared in
400 P1 Eppendorf microcentrifuge tubes and contained 65 P1 of a 1:1
mixture of the silicon oils AR 20 and.AR 200 as the middle layer and

20 #1 of 1 M.glycine containing 0.75% SDS as the bottom layer. The cells
were separated from the incubation medium and collected in the
glycine/SDS layer by centrifugation at 12,500g for 45 s in an Eppendorf
microcentrifuge. The radioactivity in 25 P1 of the incubation medium
(supernatant) was measured. The top two layers were removed by
aspiration, and this glycine/SDS layer was cut from the centrifuge tube
and.the cell pellet was resuspended in 0.5 ml H20.

14
In the first experiment using [ CIDMO, harvested cells were

resuspended to 5% (0.27 mg Chl/ml) in 25 mM Mes, pH 6.4 and a 5 ml

37

aliquot was placed in an O2 electrode chamber. The experiment was
performed as described above for the experiments analyzing 1[JG-labeled

products of Chlamdms in decreasing 0 concentrations , except that

2

after the cells were light adapted, unlabeled W was added to 1 11M,

3
and after 1 min, [140mm was added to 2.5 P1 (5.24 x 108 cpn/Pmole).
Aliquots (200 Pl) were removed throughout the aerobic (light and dark)
and anaerobic periods and centrifuged in the silicon oil tubes. In other
experiments, cells which had been resuspended in 50 m Hepes, pH 7.15,
were incubated with [1401111) (18.4 JIM) either in the light or dark or

[140mm was added to cells which were pelleted by centrifugation and

. After the

then resuspended in Hepes buffer and bubbled with N2

appropriate incubation time, aliquots were centrifuged in silicon oil
tubes as described above. Similar experiments were done with
[14C]benzoic acid which was added to 1 PM (1.1 x 108 Olin/Finale), except
that cells were resuspended in 50 at“! citrate buffer, pH 5.3.

The intracellular volume of the cell suspension was estinated as
described by Heldt (96). An 0.1 ml aliquot of the cell suspension was
incubated with 1 #1 of 31120 (500 PCi/ml) for 90 3. Following the
incubation, samples were centrifuged through the silicon oil layers as
described above. Results from this procedure gave an estimate of the
intracellular volume plus extracellular space. The volume of this
extracellular space was measured in a similar manner, except that cells

14
were incubated with 1 Pl of [ C]sorbitol (50 PCi/ml).

38
Data was analyzed by the computer program, SILOIL, written by Dr.
H. D. Husic. This program.caloulated the intracellular concentrations of
the radiolabeled.compound after correcting for the amount of the compound

in the extracellular space, and utilizing the calculated values of the

intracellular volume.

39

RESULTS AND DISCUSSION

Anaerobic Formation of Lactate
The distribution of 14C among the methanol-soluble, nonvolatile
products of Chlamdomaaas reinhazdtii was examined at decreasing 02

concentrations as monitored.in an O2 electrode (Figure 3). The 02

concentration decreased due to 02 uptake by the algae from dark
respiration.

[14C]lactate did not begin to significantly accuwlate in the cells
until the 02 concentration was less than 0.9 PM (approximately 0.1%).
The amount of lactate observed subsequently increased with time under
anaerobic conditions for at least 20 minutes. At this time, 15% of the
fixed 14C was found in lactate. In confirming experiments, aliquots of
photosynthesizing cells were removed from the light and.incubated in 1.5
ml microcentrifuge tubes in the dark with a large air space and frequent
stirring to maintain aerobic conditions for time periods of 10 s to 1 hr,
before adding methanol to terminate reactions. No [14C]lactate
accumulated aerobically in the dark (data not shown).

In other experiments, samples were removed from a suspension of
photosynthesizing cells after short times (10 s to 2 min) with [14C1HC03-

and made anaerobic by rapidly centrifuging them into a tightly packed

14
pellet in the dark. The percentage of the C in lactate reached 51%

40

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42

during centrifugation after 10 s of photosynthesis (Table 3).
Significant amounts of [14C]lactate were not observed in the cell
suspension that was killed by direct addition to methanol without prior
centrifugation. The [14C]lactate was apparently derived from the newly
labeled 14C-sugar phosphate pools because the label in this fraction
decreased.dramatically during the centrifugation period.

Although [14C]lactate was formed.in both of these short-term
experiments, the difference in the amount of [14C]lactate which
accumulated.may be explained.in part by the difference in the time period
required to Obtain anaerobic conditions. During the dark period in the

experiment using the O electrode, much of the label from the sugar

2
phosphate pool immediately flowed into TCA cycle intermediates (malate,
succinate, citrate, and fumarate) and related amino acids (glutamate,
aspartate, and glutamine). When the system became anaerobic, the sugar
phosphate pool was further depleted as lactate began to accumulate in the
cells. However, by this time in the dark, less 14C-sugar phosphates were
available to be converted to pyruvate and lactate via 3-P—glyoerate, as
45% of the original 14C fixed was already in the TCA cycle intermediates
and related amino acids. This carbon could flow back to pyruvate and
into lactate from malate by the malic enzyme reaction. Indeed, the
percentage of 14C in malate dropped slightly from 9% to 3% in the first

12 minutes of anaerobic conditions. However, this flow of carbon to

lactate appears to be slower than that from the sugar phosphates to

43

Table 3. The Effect of Killing Techniques on the 14C-I.abeled Products of
dilemmas reinhatdtfi. Cells were allgzed to photosynthesize in 3 11M
potassium phosphate buffer, pH 7.5, with [ ClNaHOO added to 1 M for

10 3. Reactions were then terminated by either adding the suspension of
cells directly to methanol or by centrifuging the cells to separate them
from the media and then resuspending them in methanol. Samples were
concentrated and analyzed by paper chromatography .

Method Used to Terminate Reactions

 

Products Methanol Centri f ugat ion ,
Directly Then Methanol
% 14C
Sugar Phosphates 74 11
D-Lactate O . 1 5 1
TCA cycle intermediates 3.6 15
Aspartate and glutamate 0.5 3.0
Alanine 0 . 9 7 . O
Glycolate, glycine, and serine 0.0 0.0

Other 21 1 3

44

lactate via 3-P-glycerate and pyruvate, as evidenced by the large
percentages of label seen in lactate in the experiment using rapid
centrifugation in which the pelleted cells were presunably anaerobic for
only 45 s before the reactions were terminated with methanol.

The anaerobic experiments performed in the O electrode or by

2
centrifugation were done with wild type Chlamydomonas cells which had

been grown photoautotrophically with 5% 00 Similar experiments were

2.

also carried out with cells grown with air levels of (Dz or in a Tris-

acetate firosphate medium in the light, and with the acetate-requiring
F-60 mutant of Chlamydomonas. In all cases, the cells produced

14C«-labeled lactate during short times under anaerobic conditions.

Additionally, all cells were shown to have levels of pyruvate reductase

1

activity (approximately 30 Pmoles NADH oxidized-hr- -mg Chl—1) comparable

to that of the wild type cells grown photoautotrophically with 5% (D2.
The partial purification and characterization of pyruvate reductase,
which catalyzes the formation of lactate in Chlamydanonas, are described
in Chapter III. The blue-green algae, Anabaena variablis and
Synechococcus leopoliensis, however, had only a trace of the NADH—
dependent pyruvate reductase activity (0 to 8 Pmoles NADH oxidized-

hr_1-mg Chl-1) . Corresponding to this, these blue-green algae did not

accunulate [14C]lactate during anaerobic conditions (data not shown).

45
Determination of the Configar_'ation of the Lactate Prodgped by

ChLagzgomonas

The lactate produced by the algae in vivo was oxidized preferentially
by the stereospecific D—lactate dehydrogenase (Table 4a). It was also
shown that the lactate formed.during assays of the pyruvate reductase
reacted preferentially with the Delectate dehydrogenase (Table 4b). The
reason for the apparent lack of absolute specificity of the
stereospecific lactate dehydrogenases for the appropriate isomer of
lactate (Table 4a) is unknown. The D—lactate used.reportedly contained
0.01%.of the L—isomer (Sigma) which does not account for the 20%
contamination observed. Also, activity (18%) was observed.with the algal
samples and Lelactate dehydrogenase. No contamination of the
Chlamydbmonas was observed.when aliquots of the culture were inoculated

on agar plates supplemented with acetate and algal medium.

Effect of Lactate Accumulation on the Intracellular pH of Chlagzgomonas
reiagardtii

Estimates of the intracellular pH of Chlamydomonas cells were
determined.by measuring the partitioning of the 14C-la'beled weak acids,
DMO and benzoic acid between the cells and medium (95-97). As observed
for the alga Chlorella saccharophila (95), the uptake of [14CIDMO by
Chlamydomonas reinhardtii did not reach equilibrium until 30 to 35 min;

4
whereas, equilibrium was reached within 2 min using [1 CJbenzoic acid.

Table 4. Determination of the Stereospecific Configuration of the
Lactate Produced by Chlamydomonas reinhandtii (a) and Formation of

D-Lactate by the Pyruvate Reductase of Chlamydomonas (b).

Sample

preparation and assays were performed as described in "Materials and

Methods".

Increase in A340

D—LDH activity

 

D-LDH L-LDH L-LDH activity
ratio
(a)
100 Fmoles D—lactate 1.062 0.260 4.08
100 Pmoles L-lactate 0.058 1.048 0.06
Algal sample 0.418 0.094 4.45
(b)
Blank (no enzyme fraction) 0 0 -
Control (no pyruvate) 0 0.018 --
Product of Pyruvate 0.150 0.033 4.54

Reductase Assay

47

Within an individual experiment, there were no significant differences in
the estimates of intracellular pH for either aerobic cells in the light
or dark, or anaerobic cells in the dark (Table 5). Intracellular
concentrations of accumulated lactate were estimated to be approximately
1 mM after 5 min of anaerObiosis. It has been previously demonstrated
that the cytoplasmic pH of a variety of unicellular algae remains
relatively constant with changing extracellular pH values ranging from
around pH 5 to 7.5 (95,97) The photosynthetic rates of Chlorella
sacchanqphila.remained.high when external pH values were varied between 3
and 9 (95). Photosynthetic rates of ChlamydomonaSWwere not impaired in
the pH range of 4.0 to 9.5, suggesting that these algae are also capable
of regulating their intracellular pH. The mechanism of this control in
algae is uncertain, but some possibilities have been critically evaluated

by Smith and.Raven (97).

48

Table 5. Estimates of the Average Intracellular pH of Chlamydomonas
reinhandtii During Aerobic (Light and Dark) and Anaerobic (Dark)
Conditions. Estimatea4of intracellxlar pH were determined by measuring
the partitioning of [ CHI!) and [ Clbenzoic acid between the cells and
the incubation medium, as described in "Materials and Methods". The
external pH in the experiment with benzoic acid was 5.29 (50 mM
citrate). The external pH in experiment 1 with DMD was 6.40 (25 mM Mes)
and in experiment 2 was 7.15 (50 mM Hepes).

[14C]Benzoic [1401M

Acid
Conditions experiment 1 experiment 2

 

 

 

estimated internal pH
Aerobic (light) 7.1 6.6 6.6
Aerobic (dark) 7.0 6.6 6.2

Anaerobic (dark) 7.2 6.6 6.3

CHAPTER III. PARTIAL PURIFICATION AND CHARACTERIZATIGI OF A PYMNATE

MASK (D-LACTATE BMW) mm W ”WI

49

50

INTRODUCTION

The accumulation of D—lactate during anaerobic conditions was
described in Chapter II. NADHedependent pyruvate reductase activity,
catalyzing the formation of D—lactate, was observed in extracts prepared
from Chlamwdbmonas. Pyruvate reductase activity has been reported for
various unicellular algae (24,32-34). under physiological conditions,
this enzyme catalyzes an essentially irreversible reaction in the
direction of pyruvate reduction; this irreversibility is also
characteristic of the cytosolic Dblactate dehydrogenase from.Escherichia
coli (55). A mitochondrial membrane-bound glycolate dehydrogenase in
Chlamwdomonas reportedly also catalyzes the oxidation of D-lactate
(36,68,69) . To relate enzyme nomenclature with the in 31' tu activity, the
soluble enzyme from Chlamydomonas will be called pyruvate reductase and
the membrane—bound activity referred to as D-lactate or glycolate
dehydrogenase. The partial purification and characterization of the
soluble pyruvate reductase from Chlamydomonas reinhardtii is reported in

this chapter.

51

MATERIALS AND METHODS

Mategials
Chlamydomonas reinhardtii UI'EX 90 were from the R. C. Starr algal

collection at the University of Texas at Austin. Cells were grown in
high phosphate medium (88) at approximately 23°C. The algae were
illuninated with 125 )AEinsteins-m-z-s“1 from cool white fluorescent
bulbs. Cultures were aerated with either air or air enriched with 5%
002.

Enzyme grade (NH4)2304 was from.the Schwartz/Mann Inc. DEAE-
cellulose (DE-52), cellulose phosphate (P-11), and carboxymethyl
cellulose (CM-23) were Obtained from Whatman Chemical Separation Ltd.
Affi—Gel Blue was from BioRad.and Sephacryl S—300 (superfine) was from
Pharmcia. Matrix gel Red A and PM-30 Diaflo ultrafiltration membranes
(43 an) were from the Amicon corporation. Chloramphenicol was from Parke-

Davis and Co. All other biochemicals, buffers, and enzymes were

purchased from Sigma Chemical Company.

Pregation of Algal Extracts

Cells were harvested as described in Chapter II. The harvested cells
were resuspended in 3 volumes (w/v) of 50 nfl potassium phosphate buffer

or 25 11M Mops buffer, pH 7.0, containing 1 mM MT and 5 mM EDTA and

52
used for the preparation of extracts. Resuspended cells were lysed by
two passes through a Yeda press (1500 psi of N2 gas for 5 min). The
solution was centrifuged at 15,000g for 15 min to remove unbroken cells
and membrane material. Pyruvate reductase activity was found in the
resulting supernatant. In an alternate procedure, the harvested cells
were resuspended.in 3 volumes (w/v) of 0.3% sodium deoxycholate or in 50
mM potassium phosphate buffer, pH 7.0, containing 3% Triton X-100, 1 mM
MT, and 5 nfi EDTA. The mixture was stirred at 4°C for 40 min. Cellular
debris was removed by centrifugation at 12,000g for 10 min and the

supernatant decanted and.used for subsequent assays. No activity was

found in the pellet fraction.

Eaayaa Assaya

MADE-dependent pyruvate reductase activity was measured.by following
the decrease in absorbance at 340 nm. Units of activity were defined as
Pmoles of NADH oxidized per min at 25°C. The 1 ml assay mixture
contained.0.1 M potassium phosphate buffer at pH 6.2 or 7.0, 0.12 mM
NADH, and 2 mM potassium pyruvate. Enzyme fraction (20 to 30 #1) was
added to initiate the reaction. Assays for hydroxypyruvate and
glyoxylate reductase activities used 2 mM hydroxypyruvate or 10 mM
glyoxylate as substrates in place of pyruvate. No NADH was oxidized in a
control cuvette containing all components except substrate. The reverse

reaction in the direction of lactate oxidation was run at pH 9.2 in 50 mM

53

sodium pyrophosphate buffer or at pH 9.4 in 25 mM Ches with 2.7 mM NAD,
enzyme fraction and 50 mM D— or Lelactate (Li salt). All assays were

performed in duplicate or triplicate and the average values are reported.

Partial Purification of the PYruvate Redaggaaa froa,Chlagxagggga§
Protein solutions were maintained at 4°C throughout the purification
stepsandlMUI'I‘and5nMEDTAwereaddedtoallbuffersusedinthe
procedures. Protein concentrations were determined.according to Lowry et
a1. (98).
Saturated (NH ) SO), adjusted to pH 7.0 with KDH, was added to the

4 2

supernatant from.the Yeda press extraction (0.67 ml (NH4)2304/ml extract)
to provide a final concentration of 40% saturation. After 30 min, the
suspension was centrifuged at 17,000g for 15 min and the precipitate
discarded. A saturated solution of (NH4)ZSO4 was slowly added to the

supernatant to 65% saturation (0.71 ml (NH4)ZSO4flml supernatant). After
60 min, the suspension was centrifuged at 17,000g for 15 min and.the
supernatant was decanted and discarded. The pellet was dissolved in 25
mM potassium phosphate buffer at pH 7.0, containing 1 mM DTT and 5 mM
EDTA. This phosphate buffer was used throughout the purification
procedures unless otherwise noted.

The dissolved pellet from the previous step was desalted on a

Sephadex G—25 column (2.5 x 12.5 cm) which was equilibrated and eluted

with the potassium phosphate buffer. Fractions containing pyruvate

S4
reductase activity eluted in the void volume from the column and were
pooled" The pooled fractions were loaded onto a column of Affi-Gel Blue,
100 to 200 mesh, (2.0 x 5.0 cm) which had been equilibrated with the
potassium phosphate buffer, pH 7.0. The column was washed with this
buffer until the A280 of the eluant was less than 0.03. The enzyme was
eluted with a linear gradient of 0 to 1 M ROI in 50 mM potassium
phosphate buffer, pH 7.0, containing 1 mM DTT and 5 mM EDTA. Pyruvate
and hydroxypyruvate reductase activities eluted.in a broad.peak with the
bulk of the protein at approximately 0.6 to 0.7 M.KC1.

The fractions containing the reductase activities were pooled and
concentrated to approximtely 25% of the original volune with a 50 ml
Amicon concentrator using a Diaflo PM—30 ultrafiltration membrane (43 m)
and 50 psi of N2 gas. The concentrated fraction was loaded onto a
Sephacryl S-300 (superfine) column (2.5 x 88 cm) which was equilibrated
and eluted with the potassium phosphate buffer, containing 0.2 M KCl.

The pyruvate reductase activity eluted.after the bulk of the protein and
before the main peak of hydroxypyruvate and glyoxylate reductase

activities. The fractions containing the pyruvate reductase activity

were pooled and used for subsequent characterization.

55

RESULTS
Partial Purification of the Ezggxate Reductase frog Chlagzgomonas

 

The steps used to partially purify the pyruvate reductase from
Chlamydomonas reinhardtii.are detailed in "Materials and Methods", and a
summary of the procedure is presented in Table 6. Before chromatography
on an Affi—Gel Blue affinity column, the pyruvate reductase activity was
not stable at 4°C. However, after elution from the dye-ligand column
with a KCl gradient, the preparation was stable and retained over 70% of
the original activity after storage at 4°C in phosphate buffer, pH 7.0,
with 0.4 to 0.6 M KCl for 1 month. Dialysis of this fraction against 25
mM potassium phosphate, pH 7.0, resulted in a 50% decrease in the
pyruvate reductase activity within a few hours, suggesting that the high
salt concentration was necessary to stabilize the pyruvate reductase.

The last step towards purification of the enzyme was gel filtration
chromatography on a Sephacryl S-300 column (Figure 4). KCl (0.2 M) was
included in the buffer, since high salt was apparently required for
enzyme stability. This step was the first in which the pyruvate
reductase activity was separated from the main peak of both the
hydroxypyruvate and glyoxylate reductase activities. Previously, Gruber
et a1. (32) concluded that separate reductases for pyruvate and

hydroxypyruvate existed in Chlorella from studies based on differences in

Table 6. Purification procedure for the pyruvate

Wanda-ans reinbandtii .

56

reductase from

 

 

Specific Activity Purification Yield
, -1
)lmoles-min -mg -fold %
. -1
protein
Yeda press extract 0.015 1.0 100
Pellet from 40 to 65%
saturated (141145304
precipitation 0 . 050 3 . 3 86
Eluate from 0-25 column 0.060 4.0 89
Affi-Gel Blue affinity
colum and concentration
through Amicon ultra-
filtration membrane PM-30 0.22 15 66
Eluate from Sephacryl
S-300 column 0.71 47 68

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enzyme stability. In Chlamydbmonas, there are also differences in the
stability of these two enzymes. For example, 50% of the original
hydroxypyruvate reductase activity in a crude extract remained after 52
hr of storage at 4°C; whereas, the pyruvate reductase activity in the
same extract declined to only 13% of the original activity.

Some hydroxypyruvate and glyoxylate reductase activity coeluted with
the pyruvate reductase activity (Figure 4). Hydroxypyruvate and
glyoxylate are substrates for L-lactate dehydrogenases from both animal
(66,80) and plant (12,72,79) sources. The algal pyruvate reductase
apparently also catalyzes the reduction of hydroxypyruvate and
glyoxylate. Hydroxypyruvate reductase, an enzyme of the oxidative
photosynthetic carbon cycle, catalyzes the MADE-dependent reduction of
hydroxypyruvate and glyoxylate, but not of pyruvate (Chapter V).

Further attempts to purify the pyruvate reductase from Chlamydomonas
have been unsuccessful. The enzyme does not quantitatively bind to DEAE-
cellulose or DEAE—Sephadex (pH 6.2 to 8.0), TEAE-cellulose (pH 7.0 to
8.0), calciumrphosphate gel (pH 6.4 or 7.0), cellulose phosphate (pH
7.0), oarboxymethyl cellulose (pH 7.0) or oxamate—agarose or red dye-
ligand affinity columns. These columns were tested at various stages
throughout the preparation. Elution of the reductase activities from the
Affi-Gel Blue column with either NADH or NADPH (1 mM) resulted in poor
recoveries, and the activity only slowly eluted from the column in a

volume of several hundred ml.

60

 

Propertie§,of the Partially Purified Ezggxate Reductase From
Chlggmghmonas

The level of pyruvate reductase activity in extracts of Chlamwdbmonas
reinhardtii (#90) grown with 5% 002 was 28 t 6 (n = 10) Pmoles NADH
oxidized-hr-l-mg cm"1 compared to the maximal Chlamydanonas
photosynthetic rate of approximately 120 to 160 Pmoles co2 fixed-hr-l-mg
Chl-1. Air—grown cells had.lower levels of activity of 18 i 6 (n = 10)

Pmoles NADH oxidizednhr-l-mg Chl-1. Pyruvate reductase activity was also
observed in other unicellular green algae, including Chlamwdbmonas
reinhardtii 137, Chlorella vulgaris, Chlorella.miniata, and DUnaliella
tertiolecta (data not shown). Pyruvate reduction was dependent on NADH;
no activity was observed.with equivalent amounts of NADPH.

A summary of some properties of the pyruvate reductase is presented
in Table 7. The pH optimum, determined for pyruvate reduction holding
the concentration of both pyruvate (2 mM) and.MADH (0.12 mM) constant,
was maximal at pH 7.0 (Figure 5). The ratio of Vmax to the apparent
Km(pyruvate) determined.at various pH values was plotted against pH (not
shown) and this was used to determine the pKe values (Table 7).
Calculations were performed as described by Segel (99) for a diprotic
system where the successive pK values of ionizable groups on the enzyme

are closer than 2.0 pH units. The pKe value of 6.4 (Table 7) possibly

indicates an imidazole group (histidine) and the value of 7.5 may

61

Table 7. Summary of’souerproperties of the pyruvate reductase from
WW reinhandtii. Estimates of pK values for ionizable groups
on the enzyme were determined as described by Segel (99). Kinetic data
was analyzed using a non-linear curve-fitting program adapted from
Duggleby (100).

 

 

pH optinun 7 0
pK 6.4
e1
pkez 7.6
pH Buffer Apparent Vmax
K (pyruvate) K
m m
m
5.7 Mes 1.4 $0.6 0.034
6.7 Mes 0.48 t 0.18 0.085
7.0 KP 0.50 t 0.10 0.083
7.4 Mops 0.93 i 0.09 0.056
7.8 Epps 1.4 $0.22 0.039
8.5 Epps 8.5 $0.7 0.005

62

Figure 5. pH Profile for the Partially Purified Pyruvate Reductase from
Chlamdomonas reinhardtii. Assays were performed as described in
"Materials and Methods" with 25 m buffers of either: Mes (A );

Mops (V); Epps (O); Ches (I).

63

 

 

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64

represent the a-amino group at the end of the polypeptide chain (99).
With hydroxypyruvate as a substrate, the pH optimum was 5.5 to 6.4.

The activity in the direction of D-lactate oxidation was tested in
the pH range between 6.2 and 9.6, and in all cases was extremely low. A
maximum rate of only 1 to 3% of the pyruvate reduction rate was observed
at pH 9.4, using 50 mM Delactate. No oxidation of L-lactate occurred.
In Chlorella, D-lactate oxidation has also been reported.to be low
compared.to the reductase reaction, and the apparent Km(D—lactate) was 40
mM (32). Apparently, little D-lactate oxidation by this enzyme occurs
under physiological conditions in algae.

The apparent Km(pyruvate) for the pyruvate reductase activity in the
crude homogenate was 1.1 mM (Figure 6b); however, after partial
purification of the enzyme, the apparent Kb value for pyruvate was 0.5 i
0.1 mM (Figure 6a). The higher apparent Km(pyruvate) for activity in
crude extracts suggests that there may be an inhibitor(s) of the enzyme
in the algal extract. Further evidence for this was that the pyruvate
reductase assay was not linear with respect to the volume of crude
extract added; activity was inhibited if more than 50 #1 of extract was
included in the assay. However, after the enzyme had been partially
purified, the assay was linear with respect to the volume of enzyme
fraction added. The apparent Km(pyruvate) of the partially purified

enzyme was relatively constant over the pH range of 5.7 to 7.8 (Table 7).

65

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The double reciprocal plot for pyruvate reductase with hydroxy-
pyruvate as the substrate yielded a biphasic plot. Estimates of the two
different apparent Rh values were approximately 46 PM and 0.3 mM. The
apparent Kh(hydroxypyruvate) for L-LDH from lettuce leaves is 3.0 mM at
pH 7.0 (72). The Kfi(hydroxypyruvate) for the NADH:hydroxypyruvate
reductase of Chlamydbmonas reinhardtii was determined to be 50 i 7 PM
(Chapter V). It is possible that the pyruvate reductase is still
contaminated.with some NADH:hydroxypyruvate reductase (see also
Figure 4).

Properties of Chlamydomanas pyruvate reductase with glyoxylate as a
sUbstrate were not determined.aince only very low levels of activity were
observed. L-lactate dehydrogenases can catalyze both the reduction of
glyoxylate to glycolate and.the oxidation of glyoxylate to oxalate
(66,101). At pH 8.5, the apparent Kh(glyoxylate) for glycolate
production by lactate dehydrogenase from lettuce leaves is 71 mM and the
Km for the glyoxylate oxidation reaction is 10.5 mM (101). Both of these
values are relatively high; hence it is doubtful that glyoxylate is a
substrate for lactate dehydrogenases under physiological conditions.

Substrate inhibition was observed at pyruvate concentrations above
2 mM for pyruvate reductase activity in crude homogenates (Figure 6b).
With the partially purified enzyme, substrate inhibition by pyruvate was
only Observed at pH values less than 7.0 and inhibition increased with

decreasing pH (Figure 7). Such pH-dependent substrate inhibition by

68

Figure 7. pH Dependence of Substrate Inhibition by Pyruvate of the
Ibmuwatelhxhxnase.

1)

(units .- ml

Activity

 

 

 

 

Figure 7.

.05 -.
pH 7.4
.04..
pH 6.7
.039 '
.oz-JL
_ pH 5.7
.01!)-
i f». é
-log [Pyruvate] (M)

70

pyruvate has been observed.for both D- and L-lactate dehydrogenases from
a number of sources, and has been attributed to the formation of an

4,
abortive ternary complex among enzyme, pyruvate and NAD (55,66,102). In

the presence of high concentrations (0.1 M) of anions including F-, Cl-,

2-

Br’, 1', so4 , scu', 0104', phosphate, and citrate, substrate inhibition
by pyruvate of the D—LDH of Polysphondvliun pallidum was at least

partially relieved (46).

Effect of Phosphate and Other Anions on Pyruvate Reductase

 

Addition of potassium phosphate to cell extracts resulted in
stimulation of the pyruvate reductase activity (Figure 8). If crude
extracts were prepared in 25 mM Mops buffer, pH 7.4, low activity was
seen in the homogenate. This activity increased with addfition of
increasing amounts of potassium phosphate up to 0.2 M, after which
inhibition was observed (data not shown). The effects of this activation
were even more dramatic at lower pH values (Figure 8). Maximum
stimulation of the pyruvate reductase was observed at pH 4.9, but the
apparent Ka(KPi) was high (48 mM) at this pH (Table 8). At pH values of
5.8 to 7.4, more physiological values for the apparent Ka(phosphate) of
approximately 4 to 11 mM were observed. However, at pH values of more
than 6.5, the stimulation by phosphate was only 1.5-fold. It is not

clear from the data presented in Table 8 whether the monovalent or

71

Figure 8. pH Profile of the Pyruvate Reductase from Cnde Extracts of
Chlamdamonas minbamltii in Both the Presence and Absence of Phosphate.
Yeda press cell extracts were prepared as described in "Materials and
Methods" using 50 mM Mops buffer, pH 7.0, containing 1 mM DHT'and.5 mM
EDTA. Assays were performed in 50 mM buffers: Acetic acid.(pH 4.9); Mes
(pH 5.8, 6.4, 6.7); Mops (pH 7.8); Epps (pH 8.5). Phosphate, when
included in the assays, was adjusted to the appropriate pH and added to

give an approximate concentration of RFC - of 5 11M per assay. Endogenous

4
phosphate was determined to be 40.2 PM Pi per assay.

 

+Ffi

72

 

 

 

 

 

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Figure 8.

7T3

Table 8. Phosphate activetion of the pyrnvete reductase frol Chlalydolooat reinhordtii.

 

laxilul Apparent 2- _ Ionic strength at

pH stiunletion [3(Kpi} Ka(P04 ) Ka(P04 ) [a Concentrations
observed of Phosphate

-fold I! I! II

4.9 77 48 0.75 47 0.063

5.8 48 6.0 0.61 5.3 0.021

6.4 6 4.2 1.4 2.8 0.025

6.5 1.7 8.0 3.1 4.9 0.052

6.7 1.5 8.0 4.0 4.0 0.02?

1.4 1.6 11 9.2 1.8 0.060

1.8 1.5 20 18 1.5 0.060

8.5 1.5 40 39 0.6 0.101

74
divalent phosphate anion (or both) is the activating species. Phosphate
did not affect the apparent Km for pyruvate (Table 7).

Other anions also stimulated the pyruvate reductase including, in the
order of decreasing effectiveness in stimulation of the apparent vmax'
Cl- > N03- > SO42- > citrate. The apparent K.a for 01- at pH 6.5 was
9 t 3 mM. Both 01' and phosphate became inhibitory if added at
concentrations greater than.2 to 3 times the apparent K8. Substituting
the cation Na+ for K+ had.no significant effect on the stimulation
observed.

Other phosphorylated.compounds only slightly stimulated the pyruvate
reductase activity: 10 mM.concentrations of fructose 1,6—P2,
3-Pbglycerate, and u-glycerol-P stimulated the enzyme 30%, 20%, and.10%,
respectively. Some L-lactate dehydrOZenases from bacterial sources are
activated.by fructose 1,6-P2 (103-105). Apparently, the binding of

fructose 1,6-P stabilizes the tetrameric form of the enzyme and

2
increases the affinity of the enzyme for pyruvate (105). On the other
hand, the L-LDH from lettuce leaves is inhibited by both fructose 1,6-P2
and 3-P-glycerate (72).

Stability of the pyruvate reductase activity was dependent on the
presence of high salt (KCl) concentrations; activity was lost if the
enzyme fraction was dialyzed against low salt buffer. It has been shown

that ions stabilize L-LDH in the tetrameric form and protect against

denaturation by guanidine and urea (66,106) Additionally, ions activate

75

the enzyme when it is in the dimeric intermediate form during
renaturation studies (106) . Most D-lactate dehydrogenases exist as

dimers (65,66) and ions may be required to stabilize the enzyme in this

conformation.
Inhibition of the 2mm Reductase by Various CM

 

ATP inhibits L-LDH from higher plants (12,72), bacteria (49) and
animal sources (66). At pH 7.0, using Km concentrations of pyruvate and
saturating NADH, the pyruvate reductase from Chlamdananas was inhibited
55% by 1 111'! ATP, and 98% by 10 nfi ATP (Table 9). In the presence of
phosphate (100 M), the inhibition by ATP was less severe. Brown et al.
(107 ) reported that phosphate partially protected against ATP inhibition
of LDH from Actinomyces viscosus. ADP and AMP were less effective
inhibitors of the pyruvate reductase.

Although oxannte inhibits both lactate dehydrogenases from a variety
of sources and the algal glycolate (D-lactate) dehydrogenase (66,77,108),
it was a poor inhibitor of the pyruvate reductase from Chlamydomonas
(Table 9), and the pyruvate reductase did not bind to oxamate- agarose.
It has been previously shown that D-lactate dehydrogenases from several
other sources do not bind to oxamate affinity columns (77) .

Two other structural analogs of pyruvate and lactate, glycolate and
glycidate, were poor inhibitors of the pyruvate reductase.

Hydroxypyridinemethylsulfonate, hydroxymethylsulfonate and other

76

Table 9. Effects of various canpounds on the pyruvate reductase fun
anlmdmnas reinhazdtii. Assays were run using K concentrations of
pyruvate (0.5 mM) and saturating concentrations of fiADH (0.12 mM) except
where indicated. Assays were performed.in either 50 mM Mes or 100 mM
potassium phosphate buffer, pH 7.0.

 

 

Compound Concentration Percent of Control
Activity

mM X
ATP 1 45

10 2
ATP + 100 mM 18 73
Krphosphate 1 57

10 36
ADPb 10 29
AMPb 10 76

b

Oxamate 10 80
Oxamate + 100 0.05 92
mM K-phosphate 1 73

10 19
Oxalate 8 71

21 9

aAssay was performed using 60 PM NADH.

b . . . .
Assay was performed with 2 mM (saturating) pyruvate

77

Table 9 (continued)

 

Cbmpound. Concentration Percent of Control
Activity

mM S

KEN 1 110

o-Phenanthroline 4 78

EDTA 10 120
b

Glycolate 20 76
, b

Gly01date 2 97

Hydroxypyridine- b 1 44

methylsulfonate
Hydroxymethgl- 1 77

sulfonate

78
sulfonates of the general structure R—CHOH-SOaNa have been shown to
inhibit enzymes which catalyze the oxidation of glycolate or lactate
(83,109). The canpound forms a complex with the product, and this
complex, in turn, inhibits the enzyme. Both of these compounds inhibited
the pyruvate reductase (Table 9) .

The D—lactate dehydrogenases of both yeast (48) and E‘uglena (35) , and
the when or-hydroxy acid dehydrogenase (64) are zinc-requiring
enzymes, and are inhibited by oxalate, EDTA, o-phenanthroline, and
cyanide. All these inhibitors reportedly interact with the essential
zinc moiety. A slight stimulation by cyanide was noted for the pyruvate
reductase activity from Chlamydomonas (Table 9). EDTA ms also slightly
stimulatory. Additionally, neither oxalate nor o—phenanthroline
significantly inhibited the pyruvate reductase activity (Table 9) . Since
none of these compounds significantly inhibited the pyruvate reductase
from Chlamydomonas, is seems unlikely that this enzyme requires zinc for
activity.

In E. 0011', the enzymes involved in the synthesis and oxidation of
D-lactate have sulfhydryl groups which must be in a reduced state for
enzyme activity to be observed (54,60). The pyruvate reductase activity
was rapidly lost in the absence of either DTT or fi-mercaptoethanol.
Gruber et a1. (32) showed that, in crude extracts prepared from Chlorella

pyrenoi close without added reducing reagents , the pyruvate reductase

79
activity decreased by greater than 90% within 90 min at 4°C. Thus, it is
prdbable that algal pyruvate reductases also have essential sulfhydryl

groups-

Evidence for the Constitutive Nature of Eyggzgte Reductase

 

D—lactate production was observed inmediately after Chlamydomonas
cells became anaerobic (Chapter II). In the rapid centrifugation
experiments, the algae were presumably anaerobic for no more than 45 s in
the pellet before they were resuspended in methanol to terminate
reactions. However, lactate had.already accumulated in these cells
(Table 3). This evidence strongly suggests that the pyruvate reductase
is present constitutively in the aerobic cells. However, during
harvesting of the algae, cells are pelleted.during centrifugation after
each wash. During this time of temporary anaerobic conditions, the
synthesis of the pyruvate reductase could have been induced.

To evaluate this possibility, Chlamydamonas cells in the log phase of
growth were incubated with either 1 mM (281 Pg/ml) cycloheximide or 2.3
mM (743 Pg/ml) Chloramphenicol for 30 min prior to harvesting. The
ChlamwdbmanaS'cells were then harvested as usual and a crude extract
prepared. In the extracts from either cycloheximide- or Chloramphenicol-
treated cells, the pyruvate reductase activity was the same as that from
untreated cells. Roessler and Lien (30) demonstrated that cycloheximide

at a level of 15 Pg/ml did enter Chlamyddmonas cells and inhibited

80
. . 14 . .
protein synthesis by greater than 98% as measured by [ C]argin1ne
incorporation into protein. Additionally, it was shown that the
induction of hydrogenase, which occurs only under anaerobic conditions,
was inhibited 70% by this level of cycloheximide, although
Chloramphenicol (500 jig/ml) was less effective (30). It was concluded

therefore, that the pyruvate reductase is constitutive in Chlamydamonas

reinhamltii .

81

DISCUSSION

Because the.pyruvate reductase is apparently constitutive in
azlwdamanas, yet D-lactate only accumulates in the algal cells during
anaerobic conditions, it is interesting to speculate on the regulation of
this enzyme in vivo. Uhder aerobic conditions, NADH might be limiting
and ATP levels high. It was shown that the enzyme was inhibited.by ATP.
Once the cells become anaerobic, ATP levels should.drop and NADH would
become more readily available for pyruvate reduction as it can no longer
be oxidized by the mitochondrial electron transport chain. Another
consideration is that during.aerobic conditions, the high Kh(pyruvate) of
the reductase may channel pyruvate towards other pathways. It is
possible, that under anaerobic conditions, pyruvate levels may begin to
increase and the pyruvate reductase would then become a significant
reaction and lead.to the observed accumulation of D—lactate. Anions, as
well as the redox state of possible essential sulfhydryl groups on the
enzyme, may also play a role in the regulation of the pyruvate reductase.

In higher plants, accumulation of L-lactate is limited by aerobic
conditions which promote NADH oxidation by mitochondria. In the light,
high levels of ATP and P-enolpyruvate may inhibit glycolysis, and have

been shown to inhibit the L—LDH of higher plants (10,12). Under

82

anaerobic conditions (i.e. in germinating seeds and roots), Lr-lactate
production can occur, subsequently causing a decrease in cellular pH
(6,11). Since low cellular pH values are undesirable, overproduction of
lactate under these conditions must be controlled. This is accomplished
in two ways (10,12). At slightly acidic pH values, NADH and ATP are
potent and cooperative inhibitors of the L-LDH, whereas at alkaline pH,
ATP only slightly inhibits the enzyme and the inhibition is competitive
with respect to NADH. Secondly, under anaerobic conditions, the decrease
in pH approaches the pH optimun of pyruvate decarboxylase which decreases
the amount of pyruvate available to be converted to lactate . Qace the
pyruvate decarboxylase is activated, the cells begin to produce ethanol
instead of lactate (10—12). Because of this control mechanism and the
ability to regenerate NAD without acid production, plants are able to
tolerate anaerobic conditions for extended time periods.

In long term (> 3 hr) anaerobic fermentation studies, D—lactate is
not normally observed as a product of starch degradation in Cblamdanonas
(18,19). It was reported in this chapter, that D—lactate, labeled with
newly fixed 14C from the sugar phosphate pools, accrmrulated in
Chlamydanonas during short time periods under anaerobic conditions. It
is suggested that during the initial minutes of anaerobiosis, reducing
equivalents (NADH) formed in the glycolytic reactions are reoxidized by

the pyruvate reductase. After initiation of starch fermentation, the

83

subsequent metabolism is apparently switched from D-lactate fermation to
mixedracid,fermentation producing formate, acetate and ethanol (18,19).
How the flow of carbon may be switched from initial Delectate formation
to the formate-producing pathway with the onset of starch breakdown is
unknown.

The decrease in cellular pH due to lactate accumulation in plant
roots is largely responsible for turning on the pathway of ethanol
fermentation. In Chlamydbmonas, however, the anaerdbic accumulation of
lactate prObably does not affect the metabolic switch to mixed acid
fermentation. No evidence for a decrease in intracellular pH was
observed with lactate accumulation in these cells (Chapter II).
Additionally, the pH optimum of the pyruvate formate—lyase is 8.0 (31),
so a decrease in cellular pH would not favor the formation of formate and
acetaldehyde from pyruvate. Rather, the activation of pyruvate formate-
lyase under anaerobic conditions is reportedly controlled by a
complicated process involving flavodoxin, S-adenosylmethionine, pyruvate

(as an allosteric effector), and a Fe-dependent activase (31).

CHAPTER IV. INHIBITIGJ OF D-lACl‘ATE AND GLleATB METAKDLISM IN A MEANT

OF W WI DEFICIENI‘ IN MIM'IONDRIAL RESPIRATION

84

85

INTRODUCTION

Despite the evidence for D—lactate accunulation in unicellular algae
(Chapter II, and refs. 14—17,23-26), the metabolic fate of lactate in the
algal cell is unknown. In this chapter, several possible fates of
D-lactate in Chlamdamonas are considered (Figure 9) . Since the soluble
pyruvate reductase is essentially irreversible under physiological
conditions as shown in Chapter III, D-lactate is probably not reoxidized
by this enzyme in viva. It is possible that D—lactate is excreted from
the algal cell into the surrounding medium, as is glycolate when it
begins to accumulate in the cell (70,78) . Alternatively, D—lactate could
be oxidized by an enzyme distinct from.the pyruvate reductase.

It has previously been reported that the algal glycolate
dehydrogenase will also catalyze the oxidation of D—lactate in vitro
(36,37,69). The algal glycolate dehydrogenase does not use 02 as the
immediate electron acceptor, but the direct electron acceptor for the
dehydrogenase is unknown (70). Both glycolate and D-lactate
dehydrogenase activities have been cytochemically localized in the

mitochondrial membrane in Chlamydomonas reinhardtii (68). The glycolate

dehydrogenase has also been localized with the mitochondrial fraction in

86

Figure 9. Possible Fates of D-Lactate in 011W minhardtii.

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88
sucrose density gradients from both unicellular green algae (34,38-40,69)
and diatoms (110,111). In the blue—green algae (cyanObacteria) glycolate
dehydrogenase has been localized with the thylakoid.membrane fraction
(112,113). In Englena gracilis it has been demonstrated that glycolate
oxidation via a membrane-bound glycolate dehydrogenase is linked.to
mitochondrial electron transport (39,40).

It is not known whether the glycolate dehydrogenase from
Chlamydbmonas or other unicellular green algae is also linked to
mitochondrial electron transport. Paul and Vblcani (69) reported.the
glycolate-dependent reduction of cytochrome c’using mitochondrial
fractions from.Ch1amydamonas. Due to the harsh conditions required to
rupture ChlamydbmonaS'cells, it has been difficult to obtain mitochondria
which possess functional, intact electron transport chains and.which are
free of chloroplast thylakoid.membranes. The presence of photosynthetic
pigments hinders attempts to study the glycolate or D-lactate dependent
reduction of the mitochondrial electron transport chain components by
difference spectra.

In this chapter, work is described in which a mutant of Chlamwdbmonas
reinhardtii deficient in cytochrome oxidase (dk97) was used to
investigate the possibility that the glycolate dehydrogenase is linked to
mitochondrial electron transport. This mutant, isolated by Wiseman et

al. (114) is unable to survive heterotrophically with acetate as a carbon

89

source in the dark, but grows as well as wild type cells photoauto-
trophically utilizing 002. The results presented in this chapter are the
first in vivo evidence demonstrating that, in unicellular algae, both
Delectate and glycolate metabolism are linked to mitochondrial

respiration.

90

MATERIALSANDMETHODS

Materials

Chlamydbmonas reinhardtii (Dang.) UTEX 90 were from the R. C. Starr
collection at the University of Texas, Austin. Chlamydomonas reinhardtii
137 was a gift from.Dr. R. K. Togasaki, Indiana University. The mutant
strains of Chlamydomonas reinbandtii deficient in mitochondrial
respiration were provided.by Dr. E. Harris at the Chlamydomonas'Culture
Center at Duke University.

[14C1NaH003 was from New England Nuclear. Potassium ferricyanide was

from the J. T. Baker Chemical Co. All other chemicals and buffers were

purchased from the Sigma Chemical Company.

Prepargtion of Algal Cell SusEnsions and Crude Homogenates

Algae were grown as described in Chapter 2 except that minimal medium
(115) was used. Cells were tested for the ability to grow in the dark
with Tris-acetate-phosphate (89) on agar plates. Cell suspensions were
prepared as described in Chapter I and were stored on ice and used within
1 hr after harvesting. Homogenates of algal cells were prepared as
described in Chapter II. The supernatant resulting from centrifugation

after cell lysis in a Yeda press was used for pyruvate reductase assays.

91

For assays of glycolate or D—lactate dehydrogenase activity, the pellet
fraction from.the Yeda pressate was resuspended in 25 mM potassium
phosphate, pH 7.0, containing 2% Triton X-100 and stirred at 4°C for 40
min. The suspension.was then centrifuged at 17,000g for 15 min, and the
resulting supernatant used for assays. For cytochrome oxidase assays,
the Yeda press pellet fraction was resuspended in 10 mM potassium

phosphate, pH 7.4, containing 1 mM EDTA and 0.1%.Tween 80.

Assays

NADflzpyruvate reductase was assayed by monitoring the decrease in
absorbence at 340 nm (Chapter III). Glycolate (Delactate) dehydrogenase
activity was assayed anaerobically, following the reduction of DCPIP at
600 nm (36). Cytochrome oxidase assays were performed as described by

Wharton and.Tzagoloff (116) measuring the oxidation of ferrocytochrome c

1 1

at 550 nm, and the extinction coefficient (550 nm) of 19.6 mM- -cm-

(117) was used.
Photosynthetic rates were measured.with a 2 or 5% suspension of
intact ChlamydbmonaS'cells in 25 mM Hepes, pH 7.5, either as the rate of

14
[ CINaHOO (lmM) incorporation or as the rate of 02 evolution in the

3

presence of 10 mM NaHCO . Dark respiration rates were measured with

3
suspensions of intact cells in a Rank Brothers (BottinghameShire) O2

electrode chamber. After equilibration to room temperature (about

92
2 min), the chamber was covered with foil and a dark cloth and rates of
02 uptake were monitored.
Glycolate was determined in samples by the Calkins method (118).

14C-I..g.beligg and Chase gmriments

 

After harvesting, suspensions of 2 or 5% (w/v) Chlamdanonas cells
were prepared in 25 mM Hepes, pH 7.5. An aliquot was placed in a 2—cm
diameter glass vial which was held in a plastic holder in a circulating
water bath at 25°C, and the contents were stirred by a magnetic bar.
After the samples were preilluninated for 2 or 3 min by 1000 PEinsteins-

-2 -1
-s

m of white light from a projector, (”ammo was added to a final

3
concentration of 1 mM. After 30 s of photosynthesis, a 200 1‘1 aliquot
was removed and added to 0.5 ml of methanol to terminate enzymatic
reactions and to solubilize the cellular contents. The retraining sample
was pipetted into a 1.5 ml microcentrifuge tube and centrifuged in a
Eppendorf microfuge for 15 s. The supernatant was aspirated with a
Pasteur pipet, and the pelleted cells were resuspended in 25 11M Hepes, pH
7.5, to the original volume of the sample. At this point, another 200 111
aliquot was removed and added to 0.5 ml of methanol as the zero-time

sample for the chase with 1112003 . NaHCO3 was added to 1 M to the

remaining cell suspension which was returned to the light. Additional
NaHOO:3 was added at 5 min intervals to maintain approximately 1 at!

concentrations. At various times up to 1 hr, 200 P1 aliquots were

removed and added to 0.5 ml methanol.

93

To assay the cellular components and.the surrounding medium
separately, aliquots were removed at various time points and cells were
immediately separated from the medium by centrifugation for 5 s in an
Eppendorf microfuge. The supernatant was removed.with a Pasteur pipet
and.added to an equal volume of methanol. The pellet was resuspended.in
methanol to a volume equal to the original volume of the aliquot.

Inhibitors were added at either 1 min into the light adaptation
period or at the beginning of the HIZCOS- chase. Glycolate solutions
used in these experiments were brought to pH 7.5 with KDH. Stock
solutions of SHAM (300 mM) were prepared.in absolute ethanol. The effect

on photosynthesis by inhibitors of the C cycle was checked by

2
preincubating the cell suspension in the light with the compound for
either 2 or 60 min. [14CJNaHOO3 was then added, and after 30 s of
photosynthesis, an equal volume of methanol was added to the cell
suspension to terminate reactions. The acidrstable radioactivity in

these samples were expressed as a percentage of the radioactivity in

control samples.

Apalysigiof 14C-habeled Products of Chlamydomggag.

Samples from 14C-labeling experiments were concentrated to
approximately 0.1 ml in a vortex evaporator and analyzed by two—
dimensional paper chromatography (92). The radioactive compounds were

detected by exposure to Kodak XAR-S X-ray film for about 2 weeks. The

94

identity of lactate and glycolate were confirmed by cochromatography with
[Mm-labeled standards. To determine the percentage of label in
compounds of interest , the areas on the chromatographs corresponding to
the spots on the X-ray film were cut out of the chromatograms and into
approximately 5 m2 pieces and placed into scintillation vials.

Compounds were eluted from the paper by adding 1.5 ml of water to each
vial, followed by gentle mixing for 2 hr. Scintillation fluid was added,
and the samples were mixed and the radioactivity counted. The
radioactivity of the compound was expressed as a percentage of the total

radioactivity (in 20 Pl) originally applied to the chromatograph.

95

RESULTS

D—Lactate Metabolism1in Chlggldbmqggg
The object of these studies was to determine the metabolic fate of
D—lactate in Chlamydcmonaswonce it accumulated anaerobically in the

cells. Initially, H12003- chase experiments were performed to study

lactate turnover in aerobic wild type cells. After photosynthesis with

[14C]NaHOO3, cells were made anaerobic by rapid.centrifugation into a

tightly packed pellet. PreviOusly, it was demonstrated.that during this
procedure, [14C]lactate accumulated.in the ChlamydbmonaS’cells (24,87).

The pelleted cells were resuspended in fresh buffer containing 1 mM

H12003-, and returned to aerobic conditions in the light. Results from a

typical chase experiment are shown in Figure 10. Aerobically, in the
light, the 14C-label was rapidly chased out of lactate in air-grown
Chlamydomonas cells. However, lactate turnover did not occur in the dark
even after 1 hr (data not shown). Rapid turnover of the 14C-label in
lactate did not occur in cells which had been grown in high (5%) 002
(Figure 10). In fact, after 1 hr, 90% of the original label in lactate
still remained (not shown). In no case was [14C]lactate observed in the

medium, indicating that D-lactate is not excreted from Chlamyddmonas

cells. Glycolate, a structural analog of lactate, is excreted from

96

Figure 10. Turnover of D—[14CJIactate in Air- and 5% (Dz—grown
Chlamdamanas. NaleCO3 chase experiments were performed as described in
"Materials and Methods" using a 5% (w/v) suspension of algae in 25 mM
Hepes, pH 7.5. Data shown are results from a typical in viva chase

experiment .

97

 

50

0/0 14c in Q - lactate

 

  

5% 002 - grown cells

 

alrbgrown cells

 

 

Figure 10.

1.0 1.5 2-0
Time of chase (min)

98

unicellular algae (70,78) when synthesized at rates faster than it can be
oxidized by glycolate dehydrogenase (119).

As shown in Chapter III, the pyruvate reductase of Chlamydomonas
probably does not catalyze the oxidation of D—lactate under physiological
conditions, so a different enzyme must catalyze lactate oxidation in
vivo. Several reports have claimed that the glycolate dehydrogenase, an
enzyme of the oxidative photosynthetic carbon cycle, catalyzes the
oxidation of both glycolate and.D—lactate in unicellular green algae
(32,36,37,38,67-69). If D—lactate is metabolized by glycolate
dehydrogenase, then one might expect that glycolate would compete for the
enzyme and possibly inhibit lactate turnover in vivo. Glycolate is
rapidly taken up by ChlamydbmonaS'cells in the light until equilibrium is
reached between the inside the cell and the surrounding medium (119).

The possibility of in vivo inhibition of D-lactate metabolism by
glycolate was investigated by performing lactate turnover experiments
with cells which had been preincubated with 10 mM K—glycolate, pH 7.5.

After photosynthesis with [14CJNaHOO , cells were pelleted by

3
centrifugation, allowing anaerobic lactate formation. The presence of
glycolate did not inhibit D-lactate production. After 1 min, the
pelleted cells were resuspended in fresh buffer, containing 10 mM
Keglyoolate. Chase experiments were then completed as described in

"Materials and Methods". In three separate trials, no inhibition of the

turnover of 14C-label in lactate was observed as compared to the controls

99

without glycolate (data not shown). These results suggest that D—lactate
and glycolate nay be oxidized by separate enzymes in Chlamydomonas. It
has been previously reported that separate dehydrogenases catalyze the

oxidation of glycolate and D-lactate in Euglena gracilis (120).

Evidggce that the Glycolate Dehydrogenase is Linked to Mitochondrial

Electron Traggpggg

 

The direct electron acceptor for either D-lactate or glycolate
oxidation is unknown. Both of these activities haye been cytochemically
localized in the mitochondrial membrane (68). Because of the location of
these activities and the fact that glycolate-dependent reduction of
cytochrome crhas been reported for Chlamydomonas, it was reasonable to
propose that the dehydrogenase(s) is linked to the mitochondrial electron
transport chain. The observation that D-lactate metabolism in
Chlamydbmonas is dependent on aerobic conditions provides further support
for this hypothesis.

Various attempts to isolate intact functional mitochondria free of
chloroplast pigments have been unsuccessful, both by us and other
researchers (114) Studies of the effects of inhibitors of mitochondrial
electron transport on glycolate and D-lactate metabolism in intact cells
are usually not feasible since either the inhibitors are impermeable to
the cells, or the inhibitors are not specific. For instance, besides

inhibiting cytochrome oxidase, cyanide also inhibits carbonic anhydrase

100

and ribulose bisphosphate carboxylase/oxygenase (and therefore, 002-
fixation) (121-123), and glycolate dehydrogenase (36,70).

Thus, we explored.the possibility of using a mutant of Chlamydbmonas
reinbardtii deficient in mitochondrial respiration to approach this
question of whether glycolate and/or D-lactate metabolism are linked to
electron transport. Several mutants originally isolated by Wiseman et
al. (114) have diminished respiratory rates and are unable to grow in the
dark on acetate. We chose the mutant designated as dk97 which is
deficient in cytochrome oxidase activity, but has approximately normal
mitochondrial ultrastructure, active cyanide—insensitive respiration, and
normal levels of other respiratoryblinked enzymes (114).

Levels of cytochrome oxidase activity were measured in detergent-
solubilized membrane fractions prepared from.Chlamydhmonaslcells by
monitoring the oxidation of ferrocytochrome 0*at 550 nm. The activity in
dk97 cells was only about 11% of that observed with wild type cells
(Table 10). Despite the decreased levels of cytochrome oxidase activity,
the mutant still exhibited dark respiration rates similar to that of wild
type cells (Table 10). However, with the mutant, the respiration rate
was inhibited 70% by 5 mM SHAM, an inhibitor of the alternative pathway
of respiration (124); SHAM alone had no significant effect on the dark
respiration rate of wild type ChlamydomonaS'cells (Table 10). Cyanide (1

mM) did not inhibit rates of respiration with the mutant more than 4%

either in the presence or absence of SHAM. In the wild type cells, the

.101

Table 10. A.comparison of mitochondrial respiration between wild type
011W reinhardtii and the dk97 mutant deficient in cytochrome

oxidase.

 

Wild Type (137)
Cells

Inutant (dk97)
Cells

 

Cytochrome Oxidase Activity

(Pmoles ferrpcytochroTe c

oxidizedm mg Chl

Dark Respiration Ra
(Pmoles O uptake hr

-1 2
ms Chl )

Inhibition of Dark
Respiration by:

5 mM SHAM

1 mM.KCN

5 mM SHAM + 1 mM KCN

93 i 18 (n = 3)

20 to 65

i 4 %
20 %
80 to 90 %

10 i 6 (n = 3)

25 to 65

7O %
i 4 %
70_to 75 %

.102
combination of cyanide and SHAM inhibited the dark respiration rate by 85
to 90%. Levels of both glycolate and D—lactate dehydrogenase activities
in extracts prepared from either dk97 cells or wild.type cells were in

1

1-mg Chi' .

the range of 3 to 4 Pmoles DCPIP reduced~hr-

Inhibition of glycolate metabolism in unicellular algae results in
increased glycolate excretion from the cells into the surrounding medium
(70,78,119,123). Such studies have been done with ChlamwdbmonaS'using
the C2 cycle inhibitors aminooxyacetate (ADA) and aminoacetonitrile
(78,87,119). Our hypothesis was that, if glycolate dehydrogenase was
linked to mitochondrial electron transport, then SHAM should cause an
increase in the rate of glycolate excretion by the mutant deficient in
cyanide-sensitive respiration. However, in SHAM-treated.wild type cells,
electrons from glycolate could still flow through the cytochrome
respiratory pathway and little or no increase in glycolate excretion
would be expected.

The results of such experiments are presented in Table 11. In the
absence of inhibitors, both wild type and.dk97 cells exhibited low rates
of glycolate excretion into the surrounding medium during
photosynthesis. The rate of glycolate excretion by wild type
ChlamydbmonaS'cells was not significantly increased in the presence of
SHAM (p > .4) Under these conditions, total glycolate excreted into the
surrounding medium ranged from 0 to 1.5 nmoles per ml of sample after

1 hr. In contrast, glycolate excretion was increased 7.4—fold with SHAM-

103

Table 11. Effect of SHAM and aminooxyacetate (ADA) on the rate of
glycolate excretion by air-grown Chlamydananas reinhardtii wild type and
dk97 cells. values are the averages i the standard deviation for the
number of experiments shown in parenthesis. The students T—test was used
to determine p values. The values for the rate of glycolate excretion in
wild type cells 1 SHAM are not significantly different (0.4 < p (0.5).
However, the differences in rates observed with dk97 cells 1 SHAM are
significantly different (p < 0.001). Additionally, rates of glycolate
excretion by SHAMetreated.dk97 cells are significantly different than the
rates observed with AOAPtreated.dk97 cells (0.01 (p < 0.025).

 

Rate of Glycolate Excretion

 

Wild type dk97
Cells Cells
-1 -1
Pmoles glycolate excreted-h -mg Chl
Control 0.03 +/- 0.07 (5) 0.46 +/- 0.46 (15)
+ 5 mM SHAM 0.12 +/- 0.27 (5) 3.6 +/- 1.8 (8)
+ 2 mM AOA 2.1 +/- 0.0 (2) 1.7 +/- 0.6 (7)

+ 5 tn”! SHAM + 2 mM AOA 2.0 +/- 0.4 (2) 3.2 +/- 0.7 (7)

104:

treated dk97 cells, and total glycolate excreted after 1 hr was 93 +/- 21
nmoles per ml sample. The rate of glycolate excretion with SHAM-treated
dk97 cells was simdlar to the estimated.maximal rates of glycolate flux
through the 02 cycle in Chlamydbmonas (119), suggesting that glycolate
oxidation was totally blocked.

Results from similar experiments with ADA-treated cells show the
expected increase of glycolate excretion by both wild type and.mutant
cells (Table 11). It was observed that the rate of excretion by dk97
cells in the presence of SHAM was higher than that observed by either the
wild type or mutant cells in the presence of ADA. It is assumed that ADA
completely blocks the transamination of glyoxylate to glycine, and that
glyoxylate is reduced to glycolate by glyoxylate reductase, since
glyoxylate is not excreted by Chlamydbmonaswcells (70,87,119). It is
therefore possible, that when the conversion of glyoxylate to glycine is
inhibited, some glyoxylate is oxidized to CO and formate (70). Under

2

normal growth conditions, photorespiratory OO2 evolution from sites other

than glycine decarboxylation is apparently negligible in both plants (70)
and Chlamydomonas (125). In higher plants, CO2 release from glyoxylate

only occurs under conditions of severe nitrogen shortages in which there

is an absence of amino donors required for the aminotransferase reactions

(126-128).

.105

SHAM (5 mM) had.no effect on the glycolate or D—lactate dehydrogenase
activity in detergent-solubilized membrane fractions. In contrast, 1 mM
cyanide inhibited these activities by 100%. Thus, in vivo inhibition of
glycolate metabolism was not due to inhibition of glycolate dehydrogenase
directly. Additionally, SHAM did not inhibit the external carbonic
anhydrase of Chlamydomonas, and SHAMetreated cells retained their high
affinity for inorganic carbon (H. D. Husic, unpublished observations).
It has previously been observed that inhibition of carbonic anhydrase
reduces the affinity of Chlamydomonas reinhardtii for inorganic carbon
and.increases glycolate excretion in Chlamydomonas (129). Because the
inorganic carbon concentrating mechanism suppresses the oxygenase
reaction of ribulose bisphosphate carboxylase/oxygenase, glycolate
synthesis is also decreased (70,119). Therefore, since the
KO.5(inorganic carbon) was not affected, the increase in glycolate
excretion observed in the presence of SHAM was not due to inhibition of
the inorganic carbon concentrating mechanism. Conversely, since SHAM-
treated dk97 cells still had.a high affinity for inorganic carbon, the
possible involvement of either glycolate metabolism or mitochondrial
respiration in the CO2 concentrating mechanism can be ruled out.

Inhibition of 14COZ-fixation was observed in SHAM-treated cells.
This inhibition increased with the length of incubation with SHAM: 10%

inhibition after 30 s, 30% inhibition after 2 min, and 50% inhibition

after 1 hr. The reason for this inhibition is unknown. It was

2106
previously demonstrated by 02 exchange studies, that SHAM did not

directly affect photosynthetic electron transport in Chlamydamonas (122).

Evidence that D—Lactate Metabolism is Linked to Mitochondrial Electron
me;

To demonstrate whether D—lactate oxidation was also linked to
mitochondrial respiration, SHAMetreated,dk97 cells were used for lactate
turnover experiments. After [14C]lactate accumulation in air-grown
cells, 5 mM SHAM was included in the resuspension buffer prior to the
chase. Results fromlsuch experiments are shown in Figure 11. In wild
type cells, [14C]lactate turned over even in the presence of SHAM, since
these cells were still able to transfer electrons to the cytochrome
pathway of the mitochondrial electron transport chain. However, lactate
metabolism was blocked in dk97 cells treated with SHAM (Figure 11).

It was also observed.from these experiments that, during the aerobic
chase in the light, [14C]glycolate accumulated in the dk97 cells treated
with SHAM (Figure 12). This confirms the experiments above; in the
presence of SHAM, glycolate metabolism is blocked in the mutant deficient
in cyanide-sensitive respiration.

Since lactate oxidation is apparently linked to mitochondrial
respiration, the possiblity existed that D-lactate is produced
aerobically in Chlamydomonas and rapidly metabolized, so that the

observed anaerobic accumulation of lactate is actually due to the

107

Figure 11. Inhibition of D—[14C]Lactate TUrnover in SHAMPtreated
arlanvdanaras dk97 Cells Deficent in Cytochrome Oxidase Activity. Chase
experiments were performed as described in "Materials and Methods". SHAM
was added to 5 mM with the resuspension buffer prior to the chase

period. An equal volume of buffer or ethanol was added to the control

cells.

 

 

 

108

 

 

3100.. d
‘N

a

(3

.‘E

‘5 so

.5

Z“

'5

.5 6°-

.2, o

a.

o \

o 40' .

Q \

‘- o\ d
°\° 20"-

k97 + SHAM

k9? - SHAMl

\.

137 + SHAM

 

1 37 SHAM

 

a £3
Time of chase (min)

Figure 11.

10

109

Figure 12. The Percent Distribution of 14C in Glycolate in W
Wild Type and dk97 Cells in the Presence and Absence of SHAM. Experiments
were performed as described for Figure 11.

 

 

 

 

C in Glycolate

14

%

10'

110

 

 

 

 

 

 

Figure 12.

dk97 + SHAM
—o
137 + SHAM
dk97 — SHAM
411g - SHAM .
5 8 5 :0

Time of chase (min)

111.

cessation of electron transport in the absence of oxygen. To test this
possibility, SHAM was added to dk97 cells during photosynthesis in the

presence of [14CJNaHCO . Aliquots of photosynthesizing cells were added

3

directly to 0.5 ml of methanol to terminate reactions without prior
centrifugation of cells into a tightly packed pellet, thereby avoiding
the introduction of anaerobic conditions. under these conditions,

significant amounts of [14C]lactate did.not accumulate even after 5 min

of labeling dk97 cells with H14003- either in the presence or absence of

SHAM (Table 12). Thus, Delectate is synthesized only during anaerobic
conditions.

As predicted from the results shown in Table 11 and Figure 12,
significant amounts of [14C]glycolate accumulated during photosynthesis

in the presence of SHAM in dk97 cells (Table 12). Relatively little

14C—labeled glycolate accumulated in the control cells.

Evidence for the Persistence of Mitochondrigl Regpiration in the Lighg

During Photosynthegis in Chlggomonas

14C-labeled intermediates of the TCA cycle, including succinate,

fumarate and m-ketoglutarate, also accumulated in the SHAM-treated mutant

cells during photosynthesis with H14CO3- (Table 12). Except for malate,

significant levels of 14C-labeled intermediates of the TCA cycle are

usually not observed during photosynthetic experiments. The data in

Table 12 indicates that when mitochondrial electron transport is

112

Table 12. The percent distribution of 14C in glycolate, TCA cycle
intermediates, and D—lzctate in air-grown Chlamydbmonasrdk97 cells after
photosynthesis with [ C]NaHOO in either the presence or absence of

5 mM SHAM. After the indicated time period of photosynthesis (either 2
or 5 min), 200 Pl aliquots of cells were added to 500 P1 of methanol to
terminate reactions. Samples were analyzed by two-dimensional paper
chromatography as described.in "Materials and Methods".

 

 

2 min photosynthesis 5 min photosynthesis
- SHAM + SHAM - SHAM + SHAM
% 14C
Glycolate 0.42 5.8 0.76 12
Malate 5.6 6.3 4.5 3.8
m-Ketoglutarate 0.40 1.8 0.20 1.7
Succinate + fumarate 0.72 2.2 0.72 2.04

D—Lactate 0.20 0.42 0.35 0.48

113
inhibited in the light, TCA cycle intermediates are no longer oxidized in
the mitochondria and begin to accumulate in the algal cell. Since these
TCA cycle intermediates are 14C--labeled, they are apparently formed from
the sugar phosphate pool which is labeled during photosynthetic 14002—

fixation. Thus, in Chlamdomonas, both the TCA cycle and mitochondrial

electron transport apparently operate in the light during photosynthesis .

114.

DISCUSSION

Evidence presented here indicates that both glycolate and D—lactate
metabolism in Chlamvdamanas are linked to the mitochondrial electron
transport chain. Glycolate-dependent 02 uptake by intact mitochondria
from Englena gracilis was also shown to be linked to electron transport
and ATP production (39,40). waever, the direct electron acceptor for
both the algal glycolate and D-lactate dehydrogenase activities is still
unknown. Glycolate dehydrogenase will not transfer electrons to various
acceptors in vitro, including 0

m, FAD, K reigns, NAD(P)+, methylene

2’ 3

blue, and nitrate (36). However, inhibition of glycolate dehydrogenase
by 2-hydroxy-3-butynoate, a specific inhibitor of flavin oxidation
reactions (130,131), suggests that a flavin may be involved in the
reaction mechanism. Although Paul and Volcani observed glycolate-
dependent cytochrome c'reduction with a mitochondrial fraction prepared
from a wall-less strain of Chlamydbmonas (69), others have reported that
the Chlamydomonas glycolate dehydrogenase does not transfer electrons to
cytochrome c in vitro (36,108).

In blue-green algae, the glycolate dehydrogenase has been localized
with the thylakoid membranes (112,113). Both the respiratory and

photosynthetic electron transport chains are also located in thylakoid

I115

membranes (131-133). Reports of the transfer of electrons from
glycolate to photosystem I in blue-green algae (134,135) could be
explained if the glycolate dehydrogenase of blue-green algae was also
linked to electron transport.

The oxidative photosynthetic carbon cycle (Figure 2) is an
energetically wasteful pathway, but the cycle may serve to utilize excess
photosynthetic energy and reducing power (70). It has been proposed that

during conditions of low 002 and/or high light intensity, the C cycle

2
may protect against photoinhibition (84,85). In higher plants, one site
in which energy is lost is in the formation of H202 during glycolate
oxidation to glyoxylate (Figure 2, reaction 3). This reaction does not
occur in unicellular algae; however, in Chlamydomonas, excess reducing
power could be utilized via the cyanide-insensitive pathway of
respiration. In higher plants, it has also been proposed that this
alternative pathway of respiration may function to drain off excess
reducing power (136).

Wiseman et al. (114) suggested that in the absence of cyanide-
insensitive respiration (i.e. in the presence of SHAM), mutants deficient
in cyanide-sensitive respiration may be lethal, because oxidized
respiratory substrates which could accept electrons from the TCA cycle
intermediates would rapidly become depleted. In mutants of photo-

synthetic organisms deficient in mitochondrial respiration, this concept

could be extended to include not only intermediates of the TCA cycle,

.116
but also substrates from the C2 cycle (glycolate and the NADH from
glycine oxidation). In the mutant, dk97, in the presence of SHAM,
electrons from glycolate can not be transferred to the mitochondrial
electron transport chain, and the cell may lose the potential for burning
excess reducing equivalents via the oxidative photosynthetic carbon
cycle. Thus, the time-dependent inhibition of photosynthesis observed in
SHAM-treated Chlamdomonas could possibly be explained by photo-
inhibition, rather than a direct effect of SHAM on some component of the
photosynthetic electron transport chain. Peltier and Thibault (122)
previously demonstrated that, in Chlamydomonas, SHAM did not directly
inhibit photosynthetic electron transport.

The accumulation of 14C-labeled intermediates of the TCA cycle during
photosynthesis by dk97 cells in the presence of SHAM provides evidence
for the operation of both the Krebs cycle and mitochondrial electron
transport in Chlamydomonas during photosynthesis. This is in agreement
with the oxygen exchange data of Peltier and Thibault (122) from which
they concluded that, in Chlamydomonas, mitochondrial respiration was not
inhibited by light. In higher plants, NADH generated during glycine
oxidation is the preferred substrate for respiration when mitochondria
are presented with a mixture of respiratory substrates (137,138).

Whether or not photorespiratory intermediates, including glycolate, also
have priority access to electron transport chain in algae remains to be

determined.

llfi7

It is uncertain whether an enzyme distinct from the glycolate
dehydrogenase catalyzes the oxidation of D-lactate in Chlamydamonas.
However, it was demonstrated that Dblactate metabolism in these cells is
linked to electron transport. Therefore, the lactate which accumulates
during anaerobic conditions in ChlamydbmonaS'can only be reoxidized to
pyruvate during aerobic conditions. It is interesting to note that
membrane-bound Delactate dehydrogenases from.both bacteria (49,59,60,61)
and yeast (48) are also linked to electron transport chains. This is in
contrast to L-lactate dehydrogenases of higher plants and.animals which

are soluble pyridine nucleotide-linked enzymes (66,71).

CHAPTER V. HDPERTIES OF NADl-IzflYDIDXYPYRINATE REDUCTASE AND

NADHJ:GLYOXYLATE REDUCTASE IN ALGAE. PARTIAL PURIFICATION AND

CHARACTERIZATION PROV] W RELVIMRDTII.

118

.119

INTRODUCTION

This chapter focuses on enzymes f ran Chlamydomonas reinhardtii which
catalyze the reduction of the m-keto acids, hydroxypyruvate and
glyoxylate, both of which are intermediates of the oxidative
photosynthetic carbon cycle (Figure 2). The specific reactions to be
discussed are shown in Figure 13. NADflzhydroxypyruvate reductase
catalyzes the reduction of both hydroxypyruvate and glyoxylate, and
there is a distinct NADPHedependent reductase which is specific for
glyoxylate. The NADH-dependent pyruvate reductase (Delactate
dehydrogenase) from ChlamydomonaS'described in Chapter III will also
catalyze the reduction of both hydroxypyruvate and glyoxylate, as do the
L-lactate dehydrogenases of plants and animals (12,66,80).

The NADH—dependent hydroxypyruvate reductase (D-glycerate
dehydrogenase) of higher plants catalyzes the interconversion between
hydroxypyruvate and D-glycerate, and also catalyzes the reduction of
glyoxylate, but not of pyruvate (139,140,141). It is well established
that the higher plant hydroxypyruvate reductase is located.in the leaf
peroxisomes (141,142). In unicellular green algae, however,

hydroxypyruvate reductase activity is associated with the mitochondrial

120

Figure 13. ix-Keto acid reductase reactions for hydroxypyruvate,
glyoxylate and pyruvate in Gilawdmas reinhardtii .
NADflzhydroxypyruvate reductase catalyzes the reduction of both
hydroxypyruvate and glyoxylate to form glycerate and glycolate,
respectively. NADPHzglyoxylate reductase specifically catalyzes the
NADPH—dependent reduction of glyoxylate. The NADlizpyruvate reductase of
Chlamydomonas catalyzes the reduction of pyruvate (to D—lactate) ,

hydroxypyruvate , and glyoxylate .

 

121

+

 

NADH NAD
(IDOOH k] C'SOOH
0:0 > HCOH
CHZOH CHZOH
HYDROXYPYRUVATE GLYCERATE

NADCPJH NADCP3+
COOH K] COOH

 

 

> I
HC=O HZCOH
GLYOXYLATE GLYCOLATE
4.
COOH ”AD” )0 (IDOOH
(lb-:0 —:=r > H?OH
CH3 CH3

PYRUVATE D-LACTATE

Figure 13.

122

fraction on sucrose gradients (34,143,144), rather than with the
peroxisomal fraction as is the case in higher plants.

Higher plants also have an NADPH:glyoxylate reductase (145) located
in the chloroplasts (141,146). The physiological role of this enzyme is
uncertain, but it may scavenge glyoxylate in the chloroplast. Glyoxylate
inhibits both ribulose bisphosphate carboxylase/oxygenase (147) and the
regeneration of its substrate, ribulose 1,5-P2 (148). In Englena,
NADPflzglyoxylate reductase activity has been found in the mitochondria,
rather than in the chloroplast (149,150). In contrast, NADPH:glyoxylate
reductase activity in the alga Chlorogonium elongatum ‘was not associated
with the mitochondrial fraction on sucrose gradients, but since
chloroplasts ruptured in these preparations, it was impossible to
determine whether the enzyme had been in the chloroplast or cytosol in
these cells (143).

Although the enzymes of the oxidative photosynthetic carbon cycle or
C2 cycle from higher plants have been isolated and well characterized
(Figure 2), relatively little work has been done with these enzymes from
algal sources (70). The presence of C2 cycle enzyme activities in

unicellular green algae has been documented, and there is sufficient

physiological evidence to demonstrate the existence of a C cycle in

2
these algae (70,151,152). Ribulose bisphosphate carboxylase/oxygenase
(153) and phosphoglycolate phosphatase (154) have been purified and

characterized from Chlamydbmonas. Although never purified from algal

123
sources, glycolate dehydrogenase has been extensively studied (36,38-
40,69,151). The glutamate/alanine:glyoxylate aminotransferase has been
purified from both E‘uglena and Chlorella (70,155). However, data on the

purification and characterization of other enzymes of the C cycle in

2
algae, including other aminotransferases, glycine decarboxylase, serine
hydroxymethyltransferase , NADH: hydroxypyruvate reductase (glycerate
dehydrogenase) , glycerate kinase , and phosphoglycerate phosphatase , is
lacking.

Furthermore, since both the NADH:hydroxypyruvate reductase and
NADszglyoxylate reductase are reportedly compartmentalized in different
organelles in algae and in higher plant leaves, it was of interest to
purify and characterize these enzymes from Chlamydomonas and to compare
the algal enzyme forms with those of higher plants. This chapter
smmarizes the partial purification and characterization of the
NADthydroxypyruvate reductase, with its associated glyoxylate reductase

activity, and a distinct NADPH—specific glyoxylate reductase from

Chlamydomonas reinhardtii .

124

MATERIALS AND METHODS

Materials

Chlamvdanonas reinbandtii (Dang.) UTEX 90, Anabaena variabilis UI'FX B
337, Synecbococcus leopoliensis UTEX 625, Chlorella vulgaris UTEX 263,
and Dimaliella tertiolecta UI‘EX LB 999 were from the R. C. Starr
collection at the University of Texas, Austin. Other Chlorella strains
were from Dr. S. Miyachi, University of Tokyo. The F-60 mutant of
Chlamydamonas was a gift from Dr. R. K. Togasaki, Indiana university, and
a yellow mutant of Chlamydomonas was isolated by Dr. B. Sears at Michigan
State university.

Enzyme grade (NH4)ZSO4 was from.the Schwarz/Mann Inc. Affi-Gel Blue
and the immuno-blot assay kit were from BioRad. Sephacryl S-300
(superfine) amd Sephadex G-200 were from Pharmacia. Diaflo PM-30
ultrafiltration membranes (43 mm) were from the Amicon corporation. All
other biochemicals, buffers, resins, and enzymes were purchased from
Sigma Chemical Company. Common laboratory chemicals were of reagent
grade and solutions were prepared in deionized, distilled.water. Anti-

barley lactate dehydrogenase serum was kindly provided by Dr. N. Hoffman

at Michigan State University.

125

Pre tion of Al Crude H enates

'Ihe unicellular green algae were grown in either minimal median (115)
or Tris-acetate-phosphate medium (89) , and the blue-green algae were
grown as described in Chapter II. Crude algal homogenates were prepared
as Yeda press extracts (Chapter III). Soluble fractions of blue-green
algae were prepared.by lysozyme treatment of the cells followed by lysis
as previously described (154). Chl concentration in algal extracts was
measured spectrophotometrically after acetone and/or methanol extraction

(91).

We

NADH-dependent hydroxypyruvate reductase was measured by following the
decrease in absorbance at 340 nm. Units of activity were Pmles NADH
oxidized per min at 25°C. The 1 ml assay mixture contained 100 m
potassium phosphate at pH 6.2 or 7.0, 0.12 mm NAD(P)H, and 2 M
hydroxypyruvate (Li salt). Enzyme fraction (20 to 30 111) was added to
initiate the reaction. Assays for the reduction of pyruvate or
glyoxylate used 2 mi“! pyruvate or 2 or 50 mm glyoxylate (Na salts),
respectively, in place of hydroxypyruvate. No NADH was oxidized in
either a control cuvette containing all components except substrate or in
the absence of enzyme sample. Reverse reactions were run at pH 9.4 in

25 mM Ches with 2.7 mm NAD, enzyme fraction and 50 m glycerate or

l 2 6
lactate . All assays were performed in at least duplicate or triplicate
and the average values are reported.
The supernatant from Yeda press extracts was also used for assays of
the C2 cycle aminotransferase reactions which were performed as described

by Nakamura and Tolbert (156) . Sample preparation and assays for

glycolate dehydrogenase were as described in Chapter IV .

Partial Purification of the Wu and Glmlate Reductases
from Chlgggomonas

Protein solutions were maintained at 4°C throughout the purification
steps. Protein concentrations were estimated according to the Lowry
procedure (98) or by measuring the absorbance of samples at 280 nm,
assuming an extinction coefficient at 280 nm of 1 mlomg.

Saturated (NH4)ZSO4, adjusted to pH 7.0 with KOH, was added to crude

extracts prepared from high (X) -grown cells (0.54 ml (NI-I4)ZSO4/ml

2

extract) to provide a final concentration of 35% saturation. After 30
min, the suspension was centrifuged at 17,000g for 15 min and the

precipitate was discarded. Solid (N114) 2804 was slowly added to the

supernatant to 70% saturation (238 mg (NH ) SO

4 2 4/m1 supernatant). After

30 to 45 min, the suspension was centrifuged at 17,000g for 15 min and

the supernatant was discarded. The pellet was dissolved in 25 mM

127

potassiun phosphate at pH 7.0, containing 1 m D'l'l‘ and 5 m EDTA. This
buffer was used throughout the purification procedure unless otherwise
noted.

To remove (NH4)ZSOI, the dissolved pellet from the previous step was
applied to a Sephadex G-25 column (2.5 x 12.5 cm) which was equilibrated
and eluted with the phosphate buffer. Fractions containing the reductase
activities eluted in the void volume from the column and were pooled.
NADPH was added to these pooled fractions to 1 mM and.the sample was then
loaded onto a 2.0 x 5.0 cm.column of Affi-Gel Blue (100 to 200 mesh)
which had been equilibrated with the phosphate buffer. The
NADPHtglyoxylate reductase did not bind to this affinity column under
these conditions. The column was then washed with the phosphate buffer
until the A.28o of the eluate was less than 0.03. The NADH-dependent
reductase activities were eluted from the affinity column with a linear
gradient of 0 to 1 M KCl in the phosphate buffer, followed by a wash with
1.5 M KCl added.to the phosphate buffer. NADH—dependent hydroxypyruvate,
glyoxylate, and pyruvate reductase activities eluted.in a broad.peak with
the bulk of the protein at approximately 0.6 to 0.7 M KCl.

The fractions eluted from the Affi-Gel Blue column containing the
NADH-dependent reductase activities were pooled and concentrated to
approximately 25% of the original volume with an Amicon concentrator

using a Diaflo PM—30 ultrafiltration membrane and 50 psi of N gas. The

2

concentrated fraction was loaded onto a Sephacryl S-300 (superfine)

.128
column (2.5 x 88 cm) which was equilibrated and eluted with the phosphate
buffer, pH 7.0 containing 0.2 M KCl. This procedure separated the
pyruvate reductase activity from.the bulk of the hydroxypyruvate and
glyoxylate reductase activities (see Figure 4).

Hydroxypyruvate reductase fractions from the S-300 column were pooled
and concentrated to approximately 6 to 8 ml with an Amicon concentrator,
again using a Diaflo PM-30 ultrafiltration.membrane and 50 to 60 psi of
N2 gas. The concentrated fraction.was loaded onto a Sephadex G-200
column (2.5 x 100 cm) which was equilibrated and eluted with 50 nM Tris-
maleate buffer, pH 6.3, containing 5 mM EDTA.and 1 mM DTT. ‘The
appropriate fractions were pooled and used for subsequent character-
ization. An equal volume of glycerol was added to the enzyme fraction

containing the hydroxypyruvate reductase which was stable as such when

stored at -20°C.

Molecular Wei t Determination of the NADH: te Reductase
The molecular weight of the NADflzhydroxypyruvate reductase was
estimated by gel filtration on a Sephadex G—200 column as described.
previously (154). A similar procedure was also performed using a
Sephadex S-300 column (2.5 x 88 cm) which was equilibrated and eluted

with the phosphate buffer, pH 7.0, containing 0.2 M KCl.

129

Mtion of Antisera Against §pinach Wte Reductase

 

Spinach hydroxypyruvate reductase (Sigma) was initially dialyzed
against 20 m Mops, pH 7.2, containing 5 ml“! EDTA and subsequently
purified by native polyacrylamide gel electrophoresis on 5% tube gels in
the absence of SDS utilizing the buffer system of Laemmli (157).
Hydroxypyruvate reductase was located by in si tu assays in native
polyacrylamide gels as described by Titus et al. (158) , and by scanning
gels monitoring the absorbance at 280 nm. The major band of
hydroxypyruvate reductase protein was cut from gels, added to 1.0 ml of
PBS (0.2 g/l KCl, 8 g/l NaCl, 0.2 g/l KHZPO4, 2
pH 7.3 with KOH) and mixed with an equal volune of complete Freud’s

0.78 g/l K HPO4, bronchi: to
adjuvant. Approximately 2 ml of the suspension containing 200 Pg of
antigen was injected subcutaneously into a farale New Zealand White
rabbit. After 28 d, the rabbit was reinjected with 100 Jig of antigen in
Freund’s incomplete adjuvant, and imlmnne serum was collected 10 (1 later.
Rabbit IgG was purified from both preimlmnne and immune serum by
loading 3 to 5 ml of serum onto a Protein A-Sepharose column (0.4 x 3.8
cm) which was equilibrated and washed with PBS. IgG was eluted from the

column with 0.1M glycine-HCl, pH 3.0, and dialyzed against PBS.

130

Inmmunodetection of Wte Reductase

Proteins from native polyacrylamide gels were transferred to
nitrocellulose as described by Titus et al. (158) . Hydroxypyruvate
reductase was probed with either a 1:200 or 1:500 dilution of the anti-
spinach hydroxypyruvate reductase serum and was detected with a goat anti-
rabbit horseradish peroxidase conjugate system, following the procedures
outlined in the inmuno-blot assay kit (BioRad) .

To determine anti-spinach hydroxypyruvate IgG birnding to either
spinach or Chlamydomonas hydroxypyruvate reductase, 0.9 ml (0.11 U) of
enzyme was incubated with 0.1 ml of either preinlnune or immune serune IgG
at 4°C for 22 h. The sample was applied to a Protein A-Sepharose column
(0.4 x 3.8 cm) which was equilibrated with PBS. Wncypyruvate
reductase activity was measured in fractions which were eluted during the
sample application. The total activity eluted was expressed as a
percentage of the total activity originally applied to the column to
determine the amount of enzyme which bound the anti-hydroxypyruvate

reductase IgG and was retained on the column.

131

RESULTS

flyggggypyguggte and Glygxylate Reductase Activitieg in Green and Blue-
mesa—Alma

The levels of hydroxypyruvate and glyoxylate reductase activities,
measured with saturating levels of substrate, in crude extracts prepared
from Chlamydamonas reinhardtii cells are shown in Table 13. These values
are compared to the maxinal photosynthetic rate ard levels of some other
C2 cycle enzymes from these cells. The levels of hydroxypyruvate
reductase observed (76 to 140 Pmoles NADH oxidizednh-l-mg Chl-1) are more
than sufficient to account for the estimated.maximal rates of carbon flow
through the C2 cycle of 5 to 15 Hmoles-h-l- mg Chl”I in Chlamydomonas
(119). Air-grown Chlamydomonas reinhardtii consistently had about 2-fold
higher levels of hydroxypyruvate reductase than found.in cells grown with
5% 002.

Similar levels of NADflzhydroxypyruvate reductase activity were also
observed in other unicellular green algae, including various strains of
Chlorella vulgaris, Chlorella miniata and Dunaliella tertiolecta
(Table 14). Glyoxylate reductase activities were also observed in these

cells (data not shown). However, only trace levels of hydroxypyruvate

reductase activity were observed in the blue-green algae, Anabaena

132

Table 13. Levels of hydroxypyruvate and glyoxylate reductase activities
in cnde extracts of Chlamdamonas reinhardtii ard a comparison to the
rate of photosynthesis and.0ther enzymes of the oxidative photosynthetic
carbon cycle in these cells. Yeda press extracts of Chlamydomonas were
prepared and assays were performed as described in "Materials and
Methods".

8The values for the phosphoglycolate phosphatase are those reported in

reference 154.
b , .
Aminotransferase assays were performed by Dr. H. D. Husnc.

CPhotosynthesis was measured either as the rate of [14C1N’aHLCO3

incorporation or the rate of 02 evolution with a 1 to 5% (20 to 100 Pg
Chl/ml) suspension of intact cells in 25 mM NaHepes, pH 7.5, provided

with 10 mM NaH003.

.133

 

Total Activity in Crude Extracts

Air-grown 5% OOZ-grown
cells cells

 

Pmoles-h-l—mg Chl-1

NADH:Hydroxypyruvate Reductase

hydroxypyruvate 140 i 20 (10) 76 i 18 (10)
glyoxylate 90 (1) 43 t 11 (6)
NADPH:Glyoxylate Reductase 23 (1) 14 i 4 (9)
Phosphoglycolate Phosphatasea 60 t 10 (4) 57 i 10 (4)

b
Glyoxylate Aminotransferases

serine:glyoxylate 33 t 10 (3) 24 i 7 (3)
alanine:glyoxylate 59 i 21 (3) 83 i 33 (3)
glutamatezglyoxylate -- 33 (1)
Glycolate Dehydrogenase 3.5 i 1.5 (6) 1.0 i 0.5 (6)
Maxigal Photosynthetic
rate 131 i 33 (5) 103 i 15 (5)

134

Table 14. NAfl-l:hydroxypyruvate reductase activities in extracts from high
(5%) (132-grown or air—grown green and blue-green algae. The activities

are expressed as the average i
determinations in parentheses .

the standard deviation for the number of

 

Species

Air-Grown 5% CD -Grown
Cells Cells

 

Chlamydamonas reinhardtii 90
Chlamdomonas reinhardtii 90
grown in the dark with

acetate as a carbon source

Chlamdomonas reinbardtii F-60
grown with acetate

Chlorella vulgaris 263
Chlorella V'ngaris C-3
Chlorella vulgaris 11-h
Chlorella miniata C-143
Dunaliella tertiolecta L8999
Anataena variabilis B337

Synechococcus leopoliensis 625

Pmoles NADH oxidized h-1 mg Chl-1
140 i 20 (10) 76 i 18 (10)
69 (1) --
69 (1) -
40 i 26 (3) 21 1 4 (3)
-- 30 i 8 (2)
-- 20 i 3 (2)
-- 32 i 2 (2)
110 (1) 50 (1)
-- 0.4 (1)
4.0 +/ 3.0 (4) 5 (1)

135

variabiliS'and.synechococcus leopoliensis. Levels of both NADH-dependent
and.MADPHédependent glyoxylate reductase activities were also low in
synechococcus leqpoliensis, ranging from 0 to 6 Pmoles NADH oxidized-
h‘l-mg Chl-1.
As with allamydamonas, air-grown cells of both Chlorella vulgaris 263
and Dunaliella tertiolecta had approximately twice the hydroxypyruvate
reductase activity found.in cells grown with 5% 002 (Table 14). In

Eremosphaem, the levels of C cycle enzymes located in the mitochondria,

2
including hydroxypyruvate reductase, glycolate dehydrogenase, serine
hydroxymethyltransferase, and serine:glyoxylate aminotransferase, were
all reportedly higher in air-grown cells than in cells which had been
grown with high 002 (34). Higher levels of glycolate dehydrogenase
activity in air—grown algae than in high (Dz—grown cells have also been
observed with other algae (70).

Levels of the hydroxypyruvate and glyoxylate reductase activities in
both wild type Cblamdamonas reinhardtii, grown in the dark with acetate
as a carbon source, and in a mutant deficient in ribulose 5-phosphate
kinase (F-60) which requires acetate for growth, were similar to those
levels observed with wild type cells grown photoautotrophically (Table
14) . Additionally, hydroxypyruvate reductase activity (expressed on a
protein basis) was similar in both a light-sensitive yellow mutant

derived from Chlamdomonas reinhandtii which was grown on acetate in

the dark and the wild type cells grown either in the dark or

.136

photoautotrophically (data not shown). Apparently, the synthesis of
hydroxypyruvate reductase in Chlamydbmonas is not light induced.as is the
case for the peroxisomal hydroxypyruvate reductase of higher plants
(142,159). The synthesis of hydroxypyruvate reductase from the alga

-Euglena is also light-independent (160).

Partial PUrification of the NADflzflygggxznggggte Reductase and.a
NADPH:Gle§zlate Reductase frqg,Chl§gzgomonas

The steps used to partially purify the hydroxypyruvate and glyxoylate
reductase activities from Chlamydamanss reinbardtii are detailed in the
"Materials and Methods", and a summary of the procedures is presented in
Table 15. Three distinct enzymes fram Chlamydbmonas were separated which
would catalyze the reduction of hydroxypyruvate and/or glyoxylate. A
NADPH-specific glyoxylate reductase was separated from the NADH-dependent
hydroxypyruvate, glyoxylate, and pyruvate reductase activities by
chromatography on an Affi-Gel Blue dye-ligand column (Figure 14).
Extracts from ChlamydomonaS‘which had been fractionated by (NH4)ZSO4
treatment and then desalted on a Sephadex G—25 column were incubated with
NADPH (added to a final concentration of 1 mM) for 10 min and.then loaded
onto the affinity column. The NADpfltglyoxylate reductase eluted during
sample application to the column, while the NADH—dependent reductase

activities bound to the column. These latter activities were eluted from

the column by a linear salt gradient of 0 to 1.0 M KCl (Figure 14).

1.3'7

Table 15. Purification procedure for the llDflzhydroxypyruvate reductase frol Chlalydononas reinhardtii.
Procedures were as described in 'Haterials and Hethods'. Data presented are results free a typical
purification.

 

Specific Activity Purificationa Yield

 

Ploleslllbfl oxidised-

Iin .nn protein -fold I

Yeda press extract

(supernatant) 0.15 1.0 100
Pellet iron 35 to 70! saturated

(834)2304 precipitation 0.29 2.0 98
Eluate frol G-ZS colunn 0.28 1.9 91
Affi-Gel Blue affinity colunn

and concentration through

Anicon ultrafiltration

nenbrane pn-so 1.4 9.9 . 52
Eluate fron Sephacryl 8-300

coluln and concentration

through Anicon neahrane PH-30 5.1 35 42
Eluate fron Sephadex G-200

coluln 18 130 36

a . . . . . . .
During the preparation of the Yeda press extract, nelhrane-hound proteins, which are a Significant
percentage of the total cellular protein, were reloved with the resulting pellet after centrifugation.
Thus, the supernatant used for further purification steps, nay already represent a Z to l-fold
purification iron the original honogenate.

138

Figure 14. Separation of’the NADPH:glyoxylate reductase fro-(the NADH-
dependent hydroxypyruvate, glyoxylate and pyruvate reductase activities
from.Ch1amydbmanas reinhardtii‘by affinity chromatography on an Affi-Gel
Blue column. [A]. The enzyme sample, following ammonium sulfate
precipitation and desalting on a Sephadex G—25 column, was incubated with
1 mM NADH-l prior to being loaded onto the affinity column. [B] The
column was washed with phosphate buffer until the A280 was less than
0.03. [C]. The other reductase activities were eluted with a 0 to 1.0 M
KCl gradient; followed by [D], a wash with 1.5 M KCl added to the
phosphate buffer. When assaying fractions, 2 m hydroxypyruvate or
pyruvate, or 2 or 50 mM glyoxylate, and 0.12 mM NADH or NADPH were used.
NADPllzglyoxylate reductase ( . ); NADH:hydroxypyruvate reductase (I);

NADHzglyoxylate reductase (V); pyruvate reductase (O) .

139

 

 

 

 

0.7 -
NADI-I'Hydmypyruwm
Reductase
0.6 -
05 _. NADH-helmets
We. .
. NADHtGlyoxykm
o4 _ R0606?“
z \
B .
v MW
£03 ._ We.
.5 .
2
0.2 -
OJ -
B i

 

48l2

Figure 14.

 

 

Fraction Number

 

1140

The NADH:hydroxypyruvate reductase was subsequently separated from
the NADH:pyruvate reductase by gel filtration on a Sephacryl S-300 column
(see Figure 4, Chapter III). The pyruvate reductase also catalyzes the
reduction of hydroxypyruvate and glyoxylate (Chapter III); as a result,
some hydroxypyruvate and glyoxylate reductase activities coelute with the
pyruvate reductase. NADH-dependent glyoxylate reductase and
NADHJhydroxypyruvate reductase activities coeluted.from the S-3OO
column. The rates of inactivation of these two activites both by heat
(60°C) and by 10 PM DTNB were identical (data not shown), suggesting that
a single enzyme catalyzes the NADH-dependent reduction of both
hydroxypyruvate and glyoxylate as does the hydroxypyruvate reductase of
higher plants (139,141).

The hydroxypyruvate reductase from Chlamydbmonas was further purified
by gel filtration on a Sephadex G—200 column (Table 15). When the
concentrated reductase fractions from the Affi-Gel Blue column were
applied directly to the 6‘200 column, poor resolution of the pyruvate and
hydroxypyruvate reductases resulted. The best resolution of these two
enzymes and the highest specific activity of the hydroxypyruvate
reductase were obtained by the procedure outlined.in Table 15 using the
two sequential gel filtration steps.

Further attempts to purify the NADflzhydroxypyruvate reductase to
homogeneity were unsuccessful. The hydroxypyruvate reductase did not

bind to ion exchange columns under a variety of conditions. Elution of

141

the enzyme from the blue—affinity column by NADH resulted in poor
recoveries and the activity only slowly eluted from the column in a
volume of several hundred.ml. A specific activity of 18 Pmoles-min-l-mg
protein."1 for the Chlamdananas hydroxypyruvate reductase following
Sephadex G-200 chromatography was considerably lower than previously
reported values of 525 Pmoles-min-l-mg protein-1 for the purified enzyme
from member cotyledons (153) and 6220 Pmolesomin-l-mg protein-1 for the
crystalline form of the enzyme from tobacco (161). On 813 polyacrylamide
gels, the hydroxypyruvate reductase (subunit molecular weight of
approximately 45 kDa) appeared to be contaminated by two major protein
bandsat 37and60kDaandtwominorbandsat 71and82kDa.

The NADpflzglyoxylate reductase which eluted from the Affi-Gel Blue
column during sample application was not stable for more than 24 hr at
4°C. At this stage of the purification, the enzyme was purified

approximately 5-fold.

Molecular—Weight of the_h_lA;DH:Hydroxypymvate Redm

The hydroxypyruvate reductase from both Chlamydamonas and spinach
(Sigma) eluted at the same position during gel filtration from either a
Sephadex G—200 or a Sephacryl S-300 column giving molecular weight
estimates of 96 kDa or 115 kDa, respectively. The molecular weight of
the spinach enzyme was previously reported to be 97 kDA (162) , and the

enzyme purified from cucumber cotyledons had a molecular weight of

142

91 to 95 kDa (158). The bacterial form of the enzyme isolated from
PSeudcmonas acidbvenans'had.a molecular weight of 72 kDa (163). All
previous reports concluded that hydroxypyruvate reductase was a dimer

composed of two identical subunits (158,162,163).

Substrate Specificity of the Reductases

The maximal rate of hydroxypyruvate reduction at pH 6.2 catalyzed.by
the hydroxypyruvate reductase of ChlamydhmonaSHwith NADPH was 30% of that
observed with NADH. The algal enzyme also reduced glyoxylate, but not
pyruvate. This distinguishes it from the algal pyruvate reductase and
lactate dehydrogenases from plants and animals which catalyze the
reduction of all three of these «eketo acids (12,24,72,80). The
hydroxypyruvate reductase from.higher plants has been shown to have
similar substrate specificity (139-141), although the bacterial enzyme
will not catalyze the reduction of glyoxylate (163). NADbdependent
glycerate oxidation was not readily catalyzed by algal hydroxypyruvate
reductase. under optimal conditions at pH values of 8.9 to 9.5 with 50
mM D— or DL-glycerate, the rate of NAD reduction was only a few percent
of the rate of hydroxypyruvate reduction. The enzyme from higher plants
catalyzes the oxidation of glycerate, but glycolate oxidation is not

observedl(141,161).

143

The NADPH:glyoxylate reductase preferentially used NADPH and did not
significantly catalyze the reduction of hydroxypyruvate or pyruvate. A
similar specificity for the NADPflzglyoxylate reductase from higher plant

chloroplasts has been observed (145).

5m Values

Some properties of the hydroxypyruvate and glyoxylate reductase
activites from Chlamydomonas'are summarized in Table 16. The apparent Km
for hydroxypyruvate of 50 PM at pH 6.2 for the enzyme from Chlamwdbmonas
is similar to values of 120 and 62 PM reported for the enzyme from
spinach (141) and cucumber cotyledons (158), respectively. The
hydroxypyruvate reductase has a relatively low affinity for glyoxylate as
demonstrated.by the high Km for the algal enzyme of 10 mM. High apparent
Kh(glyoxylate) values have also been reported for hydroxypyruvate
reductase from higher plant leaves: 15 mM, 5.7 mM, and 9 mM for the
enzymes from spinach (141), cucumber (158), and tobacco (161),
respectively. It is doubtful that such a high concentration of
glyoxylate would accumulate in the cell; therefore, it is unlikely that
the hydroxypyruvate reductase of the C2 cycle has a role in glyoxylate
reduction in viva under normal circumstances (70).

The NADpflzglyoxylate reductase from Chlamydcmonas has a much higher
affinity for glyoxylate as judged by the apparent Km(glyoxylate) of 0.10

mM, which is two orders of magnitude less the the Km(glyoxylate) of the

144

Table 16. Properties of the Hydroxypyruvate and Glyoxylate Reductase
Activities of Chlamdanonas reinhardtii. Assays were performed as in
"Materials and.Methods".

 

 

Apparent pH
K optimum
m
mM
NADH:hydroxypyruvate reductase
hydroxypyruvate 1 0.050 t 0.007 5.0 to 7.0
(at pH 6.2)
glyoxylate 9.9 i 0.5 4.5 to 8.5
(at pH 6.2)
NADPH:glyoxylate reductase 0.10 i 0.01 5.0 to 9.5
(at pH 6.2)
NADH:pyruvate reductase
pyruvate 0.5 i 0.1 7.0
(at pH 7.0)
hydroxypyruvatea 0.3, 0.046 5.5 to 6.4
(at pH 6.2)

glyoxylateb

 

aIn determinations of the affinity of the pyruvate reductase for
hydroxypyruvate, a biphasic double reciprocal plot was observed,
suggesting that the NADfizhydroxypyruvate reductase was not completely
resolved from the pyruvate reductase (see Chapter III).

b

The level of the glyoxylate reductase activity associated.with the
pyruvate reductase was extremely low; therefore, the properties of this
activity were not determined.

145

hydroxypyruvate reductase. The NADPfizglyoxylate reductases from other
sources have similar Kh values for glyoxylate: 0.13 mM from spinach

(145), 0.32 mM from tobacco (145), and 45 PM from Euglena (149).

pfl_Aptivity Profilegifor the Reductases from Chlamygpmonas

The NADH:hydroxypyruvate reductase assayed.with hydroxypyruvate and
NADH had a pH optimum of 5.0 to 7.0 when assayed using Kephosphate buffer
(Figure 15). However, a much sharper pH optimum was Observed in the
absence of phosphate (Figure 15). FUrther characterization of this
observation revealed that, in the absence of phosphate, the enzyme
exhibited substrate inhibition by hydroxypyruvate (Figure 16). The
original pH optimum studies shown in Figure 15 had.been performed using
2 mM hydroxypyruvate in each assay. At pH 6.7, in the absence of
phosphate, this concentration is above the apparent Ki value of 0.7 mM
for substrate inhibition by hydroxypyruvate (Table 17). Thus, at higher
pH values, the decreased hydroxypyruvate reductase activity was
apparently due to substrate inhibition, rather than a pH effect per se.

Phosphate appeared to partially relieve this substrate inhibition
(Figure 16). At either pH 6.2 or 7.0, phosphate did not decrease the
apparent Kh(hydroxypyruvate) as compared to values obtained in the
absence of phosphate (Table 17). However, in the presence of phosphate,

the estimated Ki(hydroxypyruvate) for substrate inhibition by

146

Figure 15. pH profile of the NAWflLydronqpymvate reductase fro-
anlamydcnanas reinbardtii. Partially purified enzyme pooled from the
Sephacryl S-3OO fractions was used for the assays which contained 2 nfl
hydroxypyruvate, 0.12 nfl NADH, 30 Pl enzyme sample, and 25 M buffer:
Acetic acid (0). Malic acid ([3), Mes (A), Mops (v), Epps (0). Class
( I ), K-phosphate (X) . All buffers were adjusted to the appropriate
pH with 10 M KOH.

 

 

147

 

 

 

 

 

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Figure 15.

148

Figure 16. Substrate inhibition of the NADl'lzhydraxypyi-uvate reductase by
hydroxypyruvate and the partial relief of this inhibition in the preserve
of phosphate.

 

1.0 ei-

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Activity (units

149

 

 

 

 

 

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' KPi. pH 6.2
4L

. . KPi. pH 7.0
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-Log Eiydroxypyruvatél (M)

Figure 16.

.150

hydroxypyruvate was increased (Table 17). Substrate inhibition for
hydroxypyruvate reductase has not been previously described.

The pH optimum of hydroxypyruvate reductase from Chlamydamonausith
glyoxylate and NADH was broad, being optimum between pH 4.5 and 8.5, and
with glyoxylate as a substrate, phosphate had no effect on the pH curve
(Figure 17). The pH optimum of the NADPflzglyoxylate reductase from
Chlamwdbmanas was dependent on the buffer used.(Figure 17). Using a
series of sulfonic acid buffers, a broad pH optimun ranging between 5.0
and 9.5 was Obtained (Figure 17).

Hydroxypyruvate reductase activity in spinach extracts had a pH
optimum of 6.0 to 6.5 when NADH was used in the reaction, and a lower pH
optimun of 5.1 if NADPH was used (141). A broader pH optimun of about
6.0 to 7.4 was observed with the spinach enzyme in the presence of
phosphate buffers (141,164). The enzyme from cucumber cotyledons had an
optimum between pH 6.9 to 7.3 with hydroxypyruvate as the substrate
(158). With glyoxylate as a substrate, the optimum for the enzyme from
both spinach (141) and cucumber cotyledons (158) was around pH 5.8 to
6.5, and was between pH 6.3 and.6.6 for the tobacco enzyme (161). The pH
optima of the NADPH-dependent glyoxylate reductases from Euglena (149)

and spinach (145) were 6.4 and 6.0 to 6.5, respectively.

151

Table 17. The effect of pH and phosphate on some kinetic paruneters of
the NADl'lzhydroxypyruvate reductase of Chlamydanonas. Estimates of
apparent Ki values were determined using a non—linear curve-fitting
program adapted from Duggleby (100). The apparent K, values for
substrate inhibition were estimated as described by bixon and Webb (165).

 

 

Apparent (a) Apparent
K for Estimated 3 K, for
Buffer pH Hydroxy— V V Hydroxy—
max max
pyruvate pyruvate
. -1
PM Pmoles-min - nfl
ml
Mes 6.1 42 i 9 0.75 0.75 ' 3.5
Mops 6.7 391 6 0.92 0.45 0.7
Mops 7.2 49! 16 0.86 0.40 --
KPi 6.2 52 i 6 1.04 1.0 21
KP 7.0 81 i 11 1.03 0.78 8.6

 

8"The ratio of observed velocity using 2 11M hydroxypyruvate to the
estimated V gives an indication of substrate inhibition. A value of
1.0 represefi no substrate inhibition; values of less than 1 indicate
that at 2 u“, the substrate hydroxypyruvate is inhibitory.

 

 

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1154

The Effect of Apion§:on Hydroxyoyruvate Reductase Activity

It has previously been Observed that the higher plant hydroxypyruvate
reductase is activated by anions in.a pHédependent manner (161,164).
NADH-dependent hydroxypyruvate reductase activity of Chlamydomanas
reinhardtii was stimulated by both chloride and phosphate anions (data
not shown). The Ka(potassium phosphate) in Mops at pH 6.7 (Mops) was
1.5 mM. As mentioned above, phosphate also partially relieved substrate
inhibition by hydroxypyruvate, but had.no apparent effect on the enzyme
when glyoxylate was the substrate. Previous work with the higher plant
enzyme showed that anions did not have the same effect on glyoxylate
reduction catalyzed.by hydroxypyruvate reductase as they did when
hydroxypyruvate was the substrate (161,164). In fact, glyoxylate
reduction by the spinach hydroxypyruvate reductase was inhibited by
several anions including, 8042-, Br-, 01-, and N03- (164). In contrast,

all of these anions stimulated.activity when hydroxypyruvate was the

substrate (164).

Immunodetection of NADH:Hydroxypyruvate Reductase

Anti-spinach hydroxypyruvate reductase serum had little or no
affinity for the enzyme from Chlamydbmonas as judged from the results of
several experiments. The anti-spinach hydroxypyruvate reductase serum
did not precipitate the Chlamydbmonas enzyme in Ouchterlony plates, and

no cross-reaction was observed on dot blots or on Western blots. In all

155

cases, there was a positive reaction between the antiserum and the
purified hydroxypyruvate reductase from spinach. When the spinach
hydroxypyruvate reductase (0.11 units) was incubated with 0.1 ml of IgG
purified from the antiserun, and then passed through a Protein A-
Sepharose column, no hydroxypyruvate reductase activity eluted,
indicating 100% binding of the IgG to the antigen. In contrast, in
identical experiments using hydroxypyruvate reductase from Chlawdanonas,
90% of the activity eluted from the Protein A-column, further indicating
that the anti-spinach hydroxypyruvate reductase IgG did not bind the
algal enzyme.

It is known that the L-lactate dehydrogenase from plant root tissue
will catalyze the reduction of hydroxypyruvate (12,72) . Anti-barley
lactate dehydrogenase did not cross react with the hydroxypyruvate

reductases from either spinach or Chlamdomonas.

.156

DISCUSSION

NADH:hydroxypyruvate reductase from either Chlamydomana510r higher
plants catalyzes the NADH-dependent reduction of both hydroxypyruvate and
glyoxylate to glycerate and glycolate, respectively. The enzyme from
either source has a high affinity for hydroxypyruvate, but has such a
high Ki for glyoxylate that glyoxylate reduction catalyzed by this enzyme
probably does not occur in vivo. The proteins from.both sources have
similar molecular weights, and the algal enzyme exhibits anion activation
for the hydroxypyruvate reduction reaction as does the enzyme from
spinach and tobacco (161,164).

Despite these similarities, some differences between the algal and
plant hydroxypyruvate reductases were noted, Antiserum prepared against
spinach hydroxypyruvate reductase did not cross-react with the algal
enzyme. The algal enzyme apparently does not catalyze the oxidation of
glycerate under physiological conditions as does the enzyme from higher
plants (70,140,166). Substrate inhibition by hydroxypyruvate was
observed with the enzyme from Chlamydbmonas, but has not been reported
for hydroxypyruvate reductase from higher plants. Furthermore, unlike
the higher plant enzyme (142,159), synthesis of the NADH:hydroxypyruvate

reductase by both Chlamydbmcnas and Englena (160) is light-independent.

1157

Another major difference between the plant and algal hydroxypyruvate
reductases is their cellular location. The plant enzyme is localized in
the peroxisomes of higher plant leaves (141,142); whereas the algal
enzyme is associated.with the mitochondrial fraction of the unicellular
green algae Chlamydbmonas (144), Chlorogonium (143) and.Eremosphaera
(34). Although these algae have peroxisomes (34,142-144), the enzymes of
the C2 cycle which are perOXisomal in leaves from higher plants, are

apparently all located in the mitochondria. This includes the glycolate i

dehydrogenase, glutamate:glyoxylate aminotransferase, serine:glyoxylate

 

aminotransferase, and.hydroxypyruvate reductase (24). Multicellular
algae resemble higher plant leaves in that, in these organisms,
hydroxypyruvate reductase and the glyoxylate aminotransferases are
located in the peroxisomes, as are glycolate oxidase and catalase
(34,70). The unicellular algae, with glycolate dehydrogenase which does
not produce H202, often have very low levels of catalase (70,142). It is
unknown why there is an evolutionary difference between the unicellular
green algae and the multicellular algae and higher plants with respect to
both the location of these photorespiratory reactions and the type of
glycolate-oxidizing enzyme present.

The NADPH-specific glyoxylate reductase of Chlamwdbmonas is also
similar to that found in higher plant chloroplasts with respect to its

substrate specificity and relatively low am for glyoxylate. It has been

suggested that this enzyme from higher plants may play a role in

£158

scavenging any glyoxylate that may enter the chloroplast since glyoxylate
inhibits ribulose bisphosphate carboxylase/oxygenase (147,148). However,
it has been reported that, in Euglena, this enzyme is located in the
mitochondria (149,150), indicating that this enzyme may have a different
function in these cells. It has been proposed that the enzyme may
function together with glycolate dehydrogenase in a glycolate/glyoxylate
shuttle or terminal oxidase system to reoxidize pyridine nucleotides
(149).

In various attempts to produce photorespiratory (C2 cycle) mutants
none has been found which is deficient in hydroxypyruvate reductase
(70,167). The reduction of hydroxypyruvate by lactate dehydrogenase may
be sufficient to metabolize hydroxypyruvate in mutants lacking the

hydroxypyruvate reductase of the C cycle. The rate of hydroxypyruvate

2
reduction catalyzed.in vitro by the pyruvate reductase (D-lactate
dehydrogenase) of Chlamydomonas appears to be sufficient for the flux of
carbon through the C2 cycle in these cells (Chapter III).

Although hydroxypyruvate and glyoxylate reductase activities were
found in all green algae tested, only trace levels of these activities
were observed in the blue—green algae Anabaena and.Synechococcus.
Additionally, glycerate kinase activity has not been detected in blue-
green algae (D. Randall, personal communication). Studies in which the

inhibitors isonicotinyl hydrazide, aminoacetonitrile, and aminooxyacetate

were used to block the C2 cycle have produced conflicting results as to

.159
whether or not these inhibitors increase glycolate excretion as they do
in the unicellular green algae (78,168-170). Results from this lab
showed that addition of ADA, which should.block the conversion of
glyoxylate to glycine, did not result in stimulation of glycolate
accumulation and excretion as it does in Chlamwdbmonas (unpublished
observations). This latter observation could be explained in part by
reduced levels of the oxygenase reaction in these cells due to the
presence of an inorganic carbon concentrating mechanism in blue-green
algae (171,172). The absence of increased glycolate excretion in the
presence of ADA might also be explained.by the extremely low levels of
glyoxylate reductase activity in these cells. This reductase is
presumably required to convert glyoxylate to glycolate, prior to
glycolate excretion, in the presence of the inhibitor. This evidence
suggests that carbon flow through the photorespiration pathway in the
blue~green algae may be low and.that there may be alterations in the
pathway as compared to other photosynthetic organisms.

At least two alternative pathways which metabolize 02 cycle
intermediates have been considered. A phosphoserine pathway, in which
3-phosphoglycerate is converted to serine, exists in some organisms
(173). Phosphoserine phosphatase and 3-P—glycerate dehydrogenase
activities were not observed in cell—free extracts of lysozyme-treated

synechococcus indicating a lack of this P-serine pathway in these cells

also (unpublished results). The tartronic semialdehyde pathway is an

160

alternative pathway of hydroxypyruvate and glyoxylate metabolism (174),
but the presence of this pathway in blue-green algae was not confirmed
(78). Thus, the metabolic pathway for glycolate, glycine, and serine in
these cells remains unknown. It is possible that small amounts of
glycine or serine produced from glycolate in the blue-green algae is all

channeled towards other biosynthetic pathways.

M AND DISCUSSIW

161

162

W AND DISCUSSION

Conclusions

D-lactate was shown to accumulate in the unicellular green alga,
Chlamydamonas reinhandtii, only under anaerobic conditions. This lactate
was 14C-la‘beledfromthe newly synthesized sugar phosphate pool
(3-P-glycerate) which becomes 1abeled.during photosynthesis with

[1401mm . Pyruvate reductase (D-lactate dehydrogenase) , which

3
catalyzes the formation of D-lactate, was partially purified and
characterized.

Both D—lactate production (14,16,17,23,25,26) and pyruvate reductase
activity have been observed in a variety of other unicellular green algae
(Table 2 and Chapter III), indicating that D-lactate metabolism may be
common to species in the phylum Chlorophycophyta. In contrast, neither
anaerobic lactate accumulation nor significant levels of pyruvate
reductase activity were observed in the blue-green algae (cyanobacteria)
synechococcus leopoliensis or Anabaena variabilis.

The oxidation of D—lactate in unicellular green algae is catalyzed by
a mitochondrial membrane-bound dehydrogenase (36-42). Whether or not

D-lactate oxidation is catalyzed by a protein distinct from glycolate

dehydrogenase, an enzyme of the oxidative photosynthetic carbon cycle,

 

163

has not been conclusively determined. Despite this, using a mutant of
ChlamwdbmonaS'deficient in.mdtochondrial respiration, evidence was
obtained which suggests that both Dblactate and glycolate metabolism are
linked.to mitochondrial electron transport (Chapter IV).

During oxidative photosynthetic carbon metabolism in green algae,
electrons from glycolate are apparently transferred to the mitochondrial
electron transport chain. In higher plants, NADH produced during glycine
oxidation (reaction 6, Figure 2) may also donate electrons to the
respiratory electron transport chain (70). However, physiological
evidence for the operation of "dark" respiration in the light is
conflicting (175), and it has been proposed that the high levels of ATP
produced by photophosphorylation should inhibit mitochondrial electron
transport (176). Studies using the mitochondrial respiratory mutant of
Chlamydcmonas provided evidence that mitochondrial respiration does occur
in the light during photosynthesis in these cells. Based on the results
of 02 exchange studies, Peltier and Thibault (122) also concluded that
mitochondrial electron transport operates in the light in Chlamydbmonas.

D-lactate metabolism in Chlamydcmonas is remarkably similar to that
in EL c011. An essentially irreversible soluble D-lactate dehydrogenase
has been purified and characterized from EC 0011 (54,55). The D-lactate
produced in the cytosol is reoxidized by a plasma membrane-bound

D-lactate dehydrogenase which is linked to the respiratory chain (59-

61). Energy produced during lactate oxidation drives the uptake of

164

growth substrates from the medium into the bacterial cell (59-61) . It is
interesting to note that Kaback and coworkers recently published work in
which they used a mutant of E. coli, deficient in one of the terminal

oxidases (cytochrome d), to study the energetics of D-lactate metabolism

via the bacterial cytochrome a pathway (177) .

Future Research

1. Mitochondrial metabolism in algae. This area of research is
relatively unexplored in several photosynthetic organisms due largely to
the difficulties encountered in obtaining intact mitochondria free of
thylakoid contamination. Sane preliminary results using a cell wall-less
mutant of Chlamydomonas, which can be ruptured by relatively gentle
procedures, and an elutriation centrifugation rotor for gradient
preparation, have been promising (R. K. Togasaki , personal
communication).

Enriched mitochondrial fractions could be used in attempts to
further purify and characterize the membrane-bound dehydrogenase(s)
catalyzing the oxidation of both D—lactate and glycolate. Kaback
(60,61,177) has studied the analogous D—lactate dehydrogenase of E. coli
in membrane vesicles prepared from plasma membrane fraction. Many of
these techniques could theoretically be applied to study respiratory-
linked dehydrogenases in algae. It is of special interest to our

laboratory to determine the direct electron acceptor for the D—lactate

 

165

and glycolate dehydrogenase reactions. If the analogy between the
bacterial and.algal D-lactate systems is complete, one would.expect the
involvement of a flavin in the transfer of electrons from.1actate (or
glycolate) to the respiratory chain.

Chlamdomonas has an active cyanide-insensitive respiratory pathway
as do at least some higher plants (136,178). The function of this
alternative pathway of respiration in algae is unknown. One possible

function, mentioned in Chapter IV, was that C cycle intermediates

2
(glycolate and glycine) could feed electrons into the cyanide insensitive
pathway in an effort to dissipate excess photosynthetic reducing power.
This could.protect the cells from photoxidation, especially during 002
shortages. Such a function has been proposed for the oxidative
photosynthetic carbon cycle of higher plants (84,85). In addition, it
has been proposed that the cyanide-insensitive pathway in higher plants
may function as an "energy overflow" systen (136). The mutants of
Chlamwdbmonaswdeficient in cytochrome oxidase activity (114) would be
useful organisms in which to study the function and possibly regulation
of the alternative pathway of respiration in algae.

2. Regulation of AnaerObic Metabolism. It is now known that
pyruvate formate-lyase is a regulatory enzyme of the mixedeacid
fermentation pathway in some green algae, including Chlamydomonas
(18,31). It is still uncertain, however, how the initial anaerobic

production of D-lactate is turned off by the cells. It has also been

.166

reported that after approximately 8 hr of anaerObiosis, Chlamydbmonas
alter their fermentative metabolism a second.time to produce ethanol and
(1)2, rather than producing formate, acetate, and ethanol (18). How and
why this metabolic change occurs is unknown.

Anaerobic metabolism.in blue-green algae has not been extensively
studied, except with respect to the hydrogenase system (22). The
pathways of anaerobic carbon metabolism are not known, but it was shown
in Chapter II that at least two blue-green algae, Synechococcus and
Anabaena, do not produce lactate anaerobically.

3. Photorespiration in Blue-green Algae. Other unanswered questions
regarding carbon metabolism in.blue-green algae are the extent to which
photorespiration occurs in these cells and the fate of glycolate produced
via the oxygenass reaction during photorespiration. The inorganic carbon
concentrating mechanism suppresses the rate of photorespiration
(70,171). Significant levels of either hydroxypyuvate reductase or
glyoxylate reductase activities were not observed in synechococcus or
Anabaena (Chapter V). Furthermore, glycerate kinase activity has not
been detected in the blue-green algae (D. Randall, personal
communication). Inhibitors of the C cycle do not cause an increase in

2

glycolate excretion in atmospheric levels of O as they do in unicellular

2
green algae (78); although increases are seen when inhibitor studies are

performed.using 100% 02 (169,170). Other possible pathways for the

metabolism of C2 cycle intermediates, including the tartronic

1167

semialdehyde pathway and the phosphoserine pathway, may also be lacking
in these organisms.

It would also be of interest to study the glycolate dehydrogenase of
blue-green algae. In these organisms, the enzyme is localized in the
thylakoid.membranes, along with both the photosynthetic and respiratory
electron transport chains (112,113,131-133). There have been reports
that glycolate could transfer electrons to photosystem I (134,135),
suggesting that the dehydrogenase is also linked.to electron transport in

these cells.

In conclusion, the metabolism of D—lactate and the C2 cycle
intermediates, glycolate, glyoxylate, and.hydroxypyruvate, has been
studied in Chlamydbmonas reinhardtii. A physiological role for aerobic
Delectate oxidation and the extent of interaction between the oxidative
photosynthetic carbon cycle (photorespiration) and.mitochondrial
respiration are yet to be determined. It is interesting to note that the
metabolism of unicellular algae is often studied as a model system for
higher plant leaves. However, it has become obvious that algae have
metabolic processes not observed in higher plants, such as a prokaryotic-
like anaerobic fermentation pathway (18,24) and an inorganic carbon
concentrating mechanism which enhances photosynthetic rates at low
concentrations of CO and suppresses the rate of photorespiration

2

(70,129). Additionally, several differences in the C2 cycle exist

.168

between the algae and.higher plants, including the type of glycolate-
oxidizing enzyme present and the location of many of the enzymes of the
cycle (70). Hence, even though 02 inhibition of photosynthesis in
Chlorella.(due to photorespiration) was observed by warburg in 1920 and
unicellular green algae were the first organisms used to elucidate the
pathways of photosynthetic carbon metabolism by Calvin and coworkers
(70), several questions regarding oxidative photosynthetic carbon

metabolism in algae remain unanswered.

APPENDIX

APPENDIX: Papers, Abstracts, and Manuscripts in Preparation.

PM:

1.

Tolbert, N. E., Husic, H. D., Husic, D. W., Moroney, J. V. and
Wilson, B. J. (1985) "Relationship of Glycolate Excretion to the
Inorganic Carbon Pool in Microalgae" In Bicarbonate Utilization of
Photosmthetic (_)_rganisms (Barry, J. A. and Lucas, W. J. eds.)
American Society of Plant Physiologists, Rockville, MD, pp. 211-227.
Husic, D. W. and Tolbert, N. E. (1985) "D—Lactate Metabolism in
Chlamdomonas reinhardtii", Plant Physiol. _7_8, 277-284.

Husic, D. W., Husic, H. D. and Tolbert, N. E. (1986) "The Oxidative
Photosynthetic Carbon Cycle or C2 Cycle", CFC Critical Reviews in
Plant Sciences, CRC Press, Boca Raton, (in press).

Husic, D. W. and Tolbert, N. E. (1986) "Properties of NADH:Hydroxy-—
pyruvate Reductase and NADPH:Glyoxylate Reductase in Algae. Partial
Purification and Characterization from Chlamydomonas reinhardtii",
Arch. Biochem. Biophys. (suhnitted)

Husic, D. W. and Tolbert, N. E. (1986) "Inhibition of Glycolate and
D—lactate Metabolism in a Mutant of Cblanydomonas reinhardtii
Deficient in Mitochondrial Respiration" (manuscript in preparation).
Husic, D. W. and Tolbert, N. E. (1986) "Inhibition of Glycolate and
D-Lactate Metabolism in a Mutant of Chlamydomonas reinhardtii
Deficient in Mitochondrial Respiration", Proc. VII Int. Congress

Photosynthesis ( in press) .

169

170

PUBLISHED ABSTRACTS:

1.

Wilson, B. J., White, D. L., and.Tolbert, N. E. (1983) "Effect of
Aminooxyacetate on Photosynthetic CO
181.

2 Fixation", Plant Physiol. zgg,
Tolbert, N. E., White, D. L., Wilson, B. J., and Reed, T. J. (1983)
"Aminooxyacetate Stimulation of Glycolate Formation and Excretion by
Algae", Plant Physiol. 12S, 182.

Husic, D. W. and Tolbert, N. E. (1984) "D-Lactate Metabolism in
Chlamydbmonas reinhardtii", Plant Physiol. 15g, 140.

Husic, D. W. and Tblbert, N. E. (1985) "Hydroxypyruvate Reductases of
Unicellular Algae", Plant Physiol. 11S, 25.

Husic, D. W. and Tolbert, N. E. (1986) "DbLactate Metabolism in the
Alga, Chlamwdbmonas reinhardtii", Fed. Proc. 45, 1891.

 

LIST OF WES

1.

10.

11.

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Storey, K. B., "A.re-evaluation of the Pasteur effect: New mechanisms
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Hochachka, P. W. and Monlnsen, T. P. (1983) "Protons and
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Hook, D. D. and.0rawford, R. M. M. (1978) Plant Life in Anaerobic
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. Drew, M. C. and Lynch, J. M. (1980) "Soil anaerobiosis,

microorganisms, and root function", Ann. Rev. Phytopathol. 18, 37-66.

. Raskin, I. and Kende, H. (1985) "Mechanism of aeration in rice",

Science ggg, 327-329.

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plants: Development of a unitary concept", Bacteriol. Rev. 25, 1-17.
Davies, D. D., Grego, S. and Kenworthy, P., (1974) "The control of
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