[1st Edition] 9781483217031

Table of contents :
Content:
Contributors to Volume 4Page ii
Front MatterPage iii
Copyright pagePage iv
List of ContributorsPage vii
PrefacePage ixBERNARD L. HORECKER, EARL R. STADTMAN
Contents of Previous VolumesPages xi-xiii
The Regulation of Arginine Metabolism in Saccharomyces cerevisiae: Exclusion Mechanisms*Pages 1-38J.M. WIAME
The Lac RepressorPages 39-75SUZANNE BOURGEOIS
L-Glutamate Dehydrogenases*Pages 77-117BARRY R. GOLDIN, CARL FRIEDEN
Regulation of Fatty Acid Biosynthesis*Pages 119-166P. ROY VAGELOS
Kinetic Analysis of Allosteric EnzymesPages 167-210KASPER KIRSCHNER
Phosphorylase and the Control of Glycogen Degradation*Pages 211-251EDMOND H. FISCHER, LUDWIG M.G. HEILMEYER JR., RICHARD H. HASCHKE
Author IndexPages 253-266
Subject IndexPages 267-268

Citation preview

Contributors to Volume 4 SUZANNE BOURGEOIS EDMOND H. FISCHER CARL FRIEDEN BARRY R. GOLDIN RICHARD H. HASCHKE LUDWIG M. G. HEILMEYER, JR. KASPER KIRSCHNER P. ROY VAGELOS J. M. WIAME

CURRENT TOPICS IN

Cellular Regulation edited by Bernard L Horecker Albert Einstein College of Medicine Bronx, New York

·

Earl R. Stadtman National Institutes of Health Bethesda, Maryland

Volume 4 1971

ACADEMIC PRESS New York and London

COPYRIGHT © 1971, BY ACADEMIC PRESS, INC. ALL RIGHTS RESERVED NO PART OF THIS BOOK MAY BE REPRODUCED IN ANY FORM, BY PHOTOSTAT, MICROFILM, RETRIEVAL SYSTEM, OR ANY OTHER MEANS, WITHOUT WRITTEN PERMISSION FROM THE PUBLISHERS.

ACADEMIC PRESS, INC. Ill Fifth Avenue, New York, New York 10003

United Kingdom Edition published by ACADEMIC PRESS, INC. (LONDON) LTD. 24/28 Oval Road, London NW1 7DD

LIBRARY OF CONGRESS CATALOG CARD NUMBER: 72-84153

PRINTED IN THE UNITED STATES OF AMERICA

List of Contributors Numbers in parentheses indicate the pages on which the authors' contributions begin.

(39), The Salk Institute for Biological Studies, San Diego, California EDMOND H. FISCHER (211), Department of Biochemistry, University of Washington, Seattle, Washington CARL FRIEDEN (77), Department of Biological Chemistry, Washington Uni­ versity School of Medicine, St. Louis, Missouri BARRY R. GOLDIN (77), Department of Biological Chemistry, Washington University of Medicine, St. Louis, Missouri RICHARD H. HASCHKE * (211), Department of Biochemistry, University of Washington, Seattle, Washington LUDWIG M. G. HEILMEYER, JR. * (211), Department of Biochemistry, Uni­ versity of Washington, Seattle, Washington KASPER KIRSCHNER (167), Department of Biochemistry, Stanford University, Stanford, California P. ROY VAGELOS (119), Department of Biological Chemistry, Washington University School of Medicine, St. Louis, Missouri J. M. WIAME (1), Laboratoire de Microbiologie de V Université de Bruxelles and Institut de Recherches du CE.R.I.A., Bruxelles, Belgium SUZANNE BOURGEOIS

* Present address: Department of Physiological Chemistry, University of Würzburg, Würzburg, Germany. vu

Preface Recent years have witnessed rapid advances in our knowledge of the basic mechanisms involved in the regulation of diverse cellular activities such as intermediary metabolism, the transfer of genetic information, membrane permeability, and cellular differentiation and other organ functions. Information gained from the detailed analyses of a large num­ ber of isolated enzyme systems, together with results derived from physio­ logical investigations of metabolic processes in vivo, constitutes an everincreasing body of knowledge from which important generalized concepts and basic principles of cellular regulation are beginning to emerge. How­ ever, so rapid are the present advances in the general area of cellular regulation and so diverse are the disciplines involved, that it has become a formidable task for even the expert in a specialized area to keep abreast of the progress in his field. This series of volumes is concerned with such recent developments in various areas of cellular regulation. We do not in­ tend that it will consist of comprehensive annual reviews of the literature. We hope rather that it wTill constitute a medium which will, on the one hand, provide contributing authors with an opportunity to summarize progress in specialized areas of study that have undergone substantial de­ velopments and, on the other hand, serve as a forum for the enunciation of general principles and for the formulation of provocative theories and novel concepts. To this end editorial review of individual contributions will be concerned primarily with the clarity of presentation and con­ formity to publication policies. It is hoped in this manner to bring together current knowledge of various aspects of cellular regulation so as both to enlighten the uninformed and to provide a base of knowledge for those engaged in research in this subject. BERNARD L. HORECKER EARL R. STADTMAN

IX

Contents of Previous Volumes Volume 1 Conformational Aspects of Enzyme Regulation D. E. Koshland, Jr. Limitation of Metabolite Concentrations and the Conservation of Solvent Capacity in the Living Cell Daniel E. Atkinson The Role of Equilibria in the Regulation of Metabolism H. A. Krebs Regulation of the Biosynthesis of the Branched-Chain Amino Acids H. E. Umbarger On the Roles of Synthesis and Degradation in Regulation of Enzyme Levels in Mammalian Tissues Robert T. Schimke The Regulation of the Biosynthesis of a-1,4-Glucans in Bacteria and Plants Jack Preiss Allosteric L-Threonine Dehydrases of Microorganisms W. A. Wood The Aspartokinases and Homoserine Dehydrogenases of Escherichia coli Georges N. Cohen Pyruvate Dehydrogenase Complex Lester J. Reed xi

XÜ

CONTENTS OF PREVIOUS VOLUMES

Pyruvate Carboxylase Merton F. Utter and Michael C. Scrutton Author Index—Subject Index

Volume 2 DPN-Linked Isocitrate Dehydrogenase of Animal Tissues Gerhard W. E. Plant The Regulation of Biosynthesis of Aromatic Amino Acids and Vitamins J. Pittard and F. Gibson Regulation of Cholesterol Biosynthesis in Normal and Malignant Tissues Marvin D. Siperstein The Biogenesis of Yeast Mitochondria Anthony W. Linnane and J. M. Haslant Fructose 1,6-Diphosphatase from Rabbit Liver S. Pontremoli and B. L. Horecker The Role of Phosphoribosyltransferases in Purine Metabolism Kart 0. Raivio and J. Edwin Seegmiller Concentrations of Metabolites and Binding Sites. Implications in Metabolic Regulation A. Sols and R. Marco A Discussion of the Regulatory Properties of Aspartate Transcarbamylase from Escherichia coli J. C. Gerhart Author Index—Subject Index

CONTENTS OF PREVIOUS VOLUMES

Volume 3 The Regulation of Branched and Converging Pathways B. D. Sanwal, M. Kapoor, and H. Duckworth The Role of Cyclic AMP in Bacteria Robert L. Penman and Ira Pastan Cell Surfaces in Neoplastic Transformation Max M. Burger Glycogen Synthase and Its Control Joseph Lamer and Carlos Villar-Palasi The Regulation of Pyruvate Kinase Werner Seubert and Wilhelm Schoner Author Index—Subject Index

The Regulation of Arginine Metabolism in Saccharomyces cerevisiae: Exclusion Mechanisms I

J . M . WlAME

I I I

Laboratoire de Microbiologie de V Université de Bruxelles and Institut de Recherches du C.E.R.I.A.,Bruxelles, Belgium

I. Introduction I I . Inhibition of Ornithine Transcarbamylase by Epiarginase A. In Situ Suppression of Ornithine Transcarbamylase Activity by a Specific Binding Protein (Epiprotein) B. Suppression in Cell-Free Extracts and the Participation of Effectors C. The Nature of the Epiprotein and I t s Binding with Ornithine Transcarbamylase D. Physiological Significance E. Specificity of Binding F. The Regulatory Site of Ornithine Transcarbamylase I I I . Common Genetic Elements in the Control of Biosynthetic and Degradative Enzyme Synthesis A. Reciprocal Balance between Biosynthetic and Degradative Enzyme Syntheses B. Negative Type of Control of Degradative Enzymes C. Nitrogen Catabolite Repression IV. Conclusions A. Concurrent Pathways B. Epienzymatic Control C. Other Considerations References

1 4 4 6 10 13 14 15 21 21 24 32 33 33 34 36 37

I. Introduction Arginine biosynthesis and degradation in Saccharomyces cerevisiae are represented in a simplified form in Fig. 1. A detailed scheme and summary of the regulation of the whole pathways with references is given else­ where {45) * S. cerevisiae is able to make its own arginine from ammonium as sole nitrogen source. This implies the transfer of NH 4 + into glutamate which is transformed into ornithine; NH 4 + is also transferred into glutamine which is the donor of NH 3 for carbamyl phosphate synthesis. Aspartate is the * Unpublished data reported in this publication have been supported by a grant from the "Fonds de la Recherche Fondamentale Collective." 1

2

J. M. WIAME

NH 3

t

Glutamate

M IDehydrlrogenasel f ate Glutamate

^

NH3

Urea

I Ornithine I f Itransaminasel M Ornithine

lì JL.

^ ^ % Arginine

û

\ > _ ^ T^,,^,^"0'^** ATP Aspartate

Glutamate" ATP NH3 .

> Gì utamine 2ATP + C02

I Carbamyl phosphateι

Pyrimidine

FIG. 1. Simplified scheme of arginine metabolism in Saccharomyces cerevisiae. => anabolic reaction; ^ catabolic reaction. OTCase, ornithine transcarbamylase.

donor of the fourth nitrogen atom of the arginine molecule. We shall be concerned here with the activity of ornithine transcarbamylase, which catalyzes the condensation of carbamyl phosphate with ornithine, giving citrulline, a precursor of arginine. Arginine can also be used as the only nitrogen nutrient; under these conditions, arginine is degraded by arginase into ornithine and urea, the δ-nitrogen of ornithine is transferred to glu­ tamate by δ-L-ornithine transaminase, and the «-nitrogen appears also in glutamate by the dehydrogenation of the pyrroline carboxylate resulting from the cyclization of glutamic semialdehyde, the second product of transamination. Biosynthesis and degradation offer a typical case of concurrent pathways which have to be regulated in opposite way. Recip­ rocal exclusion of the two metabolisms can be exerted both at the level of enzyme activity and at the level of enzyme synthesis. Table I illustrates one aspect of this exclusion by considering the activity of a typical biosynthetic enzyme, ornithine transcarbamylase, and a typical catabolic enzyme, arginase, under four conditions of growth of S. cerevisiae. This shows the usual response for regulation of enzyme synthesis: the presence of end products strongly represses the synthesis of the biosynthetic enzyme. The synthesis of the catabolic enzyme is under two controls: the presence of arginine exerts an induction, but the full degradative capacity occurs only when ammonium, the cheapest and best source for nitrogen, is no longer available, i.e., when the cataboliti repression (21, 32) is suppressed.

3

A R G I N I N E METABOLISM I N SACCHAROMYCES

In the present case, one may speak of a nitrogen catabolito repression, since arginine alone cannot be used as a carbon source. All the work presented here is done with glucose as carbon source for growth (8). In principle, exclusion of two concurrent pathways could have been reached by distinct and independent mechanisms. Distinct repressors with appropriate affinity for effectors might have governed the synthesis of the enzymes by the classical mechanisms described in bacteria (19, 42). At the level of enzyme activity, appropriate quaternary structures and affinity for effectors and substrate might have produced the desired result (81 ). Typical examples can be found in the study of the biosynthetic and the degradative threonine deaminases (41, 43, 46) and in the case of the biosynthetic and the degradative ornithine transcarbamylases of Pseudomonas (16, 33, 88, 44). In the case of S. cerevisiae, additional mechanisms appear to function both at the level of the control of enzyme activity (Section II) and at the level of enzyme synthesis (Section III). In both cases a genetically deter­ mined element, either an enzyme or a repressor, acts simultaneously to TABLE I REPRESSION, INDUCTION, AND NITROGEN CATABOLITE REPRESSION ILLUSTRATED IN THE ARGININE METABOLISM IN Saccharomyces cerevisiae?1

Nitrogen nutrients available NH4+

Generation time (minutes at 29°C) 120

Enzyme activities (/xmoles product formed, mg protein -1 , hour -1 , at 30°C) Pool of arginine (IO -3 M) 18

L-Ornithine transcarbamylase 30N repression

N H 4 + + L-arginine

120

72

2,

repression

L-Arginine

150

190

NH 4 + limiting in chemostat

600

—

a

1 —

Data from Ramos et al. {34) and this paper, Table III, p. 22.

Arginase

4

J. M. WIAME

control the two concurrent activities. The need for additional devices for the control of concurrent reactions has been foreseen by Stachow and Sanwal (87), when studying the regulation of the two glutamic dehydrogenases of Neurospora. It is obvious that such mechanisms occur because of a peculiar metabolic situation. In addition, they may be an expression of a refined evolution to be found in eukaryotes and not in prokaryotes. This remains to be investi­ gated. II. Inhibition of Ornithine Transcarbamylase by Epiarginase A. In Situ Suppression of Ornithine Transcarbamylase Activity by a Specific Binding Protein (Epiprotein) (4) The first indications of an unusual mechanism of regulation of ornithine transcarbamylase (OTCase) activity were obtained during a comparison of the OTCase activity measured in nystatin-treated cells with the activity of this enzyme in cell-free extracts. With the last method, the differential rate of increase in activity follows the classical behavior: addition of arginine into minimal medium (NEU* as nitrogen source) stops further increase in OTCase synthesis almost completely. If the activity is measured in samples rendered permeable by treatment with nystatin, the evolution of activity, which is normal in minimal medium, becomes abnormal after addition of arginine: there is a rapid loss of activity (Fig. 2). The loss of activity is due to the suppression of OTCase. If the nystatin-treated cells are crushed in a French press, the activity is recovered. The suppression can also be removed in "permeabilized" cells by 5-minute heating at

I

1

0.6

I

I

OD

I

1.0

FIG. 2. Suppression of ornithine transcarbamylase (OTCase) activity tested in cells rendered permeable by treatment with mystatin. OTCase activity during shift from minimum to minimum + arginine with a repressible strain (1705d). Arginine is added at the times shown by the arrows (4).

ARGININE METABOLISM IN SACCHAROMYCES

O

40

> *>20 σ

o

\o w

30

40

50

70#C

60

FIG. 3. Thermoreactivation (5 minutes) of ornithine transcarbamylase (OTCase) activity of "permeabilized" cells before ( O—O) and after ( # — # ) addition of arginine for 1 hour. An exponentially growing culture is divided into two parts: one is cooled; to the other arginine is added for 1 hour as in the experiment described in Fig. 2. The two types of suspensions are "permeabilized," and samples are heated for 5 minutes at different temperatures. After cooling, the OTCase activity is measured (4). 59°C (Fig. 3). Suppression disappears slowly when the cells are kept at 0° at neutral pH, and quickly at pH 5. The nature of the suppressing agent and of its mechanism of action is indicated by the experiment reported in Fig. 4. No suppression occurs if cycloheximide is added at the same time as arginine. The process appears to be dependent on protein synthesis. If cycloheximide is added during the process of suppression, no further inhibition occurs. This shows that the

Id) 120 -

Arg λ

>

o σ

Actidione at (1)

\Jl2) &—·—

10

Actidione at (2)

σ o

£

1

0.6

0.8 OD

1

1

1.0

FIG. 4. Action of Actidione on the inactivation of ornithine transcarbamylase (OTCase). Arginine added at the time shown by the arrow. The solid line indicates the evolution of the activity as shown in Fig. 2. Actidione is added to two parts of this culture at (1) and at (2). The growth stops and the broken lines indicate the enzyme activity, which for the sake of comparison is plotted on the same ordinate as the sample of the growing culture taken at the same time (4), although in this case there is no growth.

6

J. M. WIAME

150

-fi 00 T3 σ

Q>

8 50 O 0

1.0 OD

20

FIG. 5. Ornithine transcarbamylase (OTCase) activity during shift from minimum to minimum + arginine with a nonrepressible strain (1708c). Arginine is added at the time shown by the arrow (4) ·

protein responsible for suppression does not act by a catalytic process, and suggests that it acts by stoichiometric binding. For that reason, we pro­ posed the term epiprotein for this protein. The suppression was also inhibited, but more slowly, by growth in the presence of thienylalanine. As the process was observed in situ, there is little doubt as to its physio­ logical role. Some time before this finding, a regulatory mutation (argR) was found {2) which provokes derepression and constitutivity of OTCase and of some other anabolic enzymes. In this mutant, the suppression does not occur (Fig. 5). This served as an indication that the suppression represents a form of regulation which acts in addition to the classical repression and feedback inhibition observed in Saccharomyces cerevisiae (1, 10). B. Suppression in Cell-Free Extracts and the Participation of Effectors {25-28) The fact that suppression occurs in situ and is released in cell-free extracts, suggests that one needs to work at high concentration of protein to reproduce the effect. For this reason, enzyme activity was measured with short time incubation, and the usual assay temperature of 30° was reduced to 15°. The suppression could be observed by adding a crude extract of yeast expected to contain the epiprotein (such as an extract from cells grown on arginine) to a crude extract of OTCase (from cells grown in minimal medium) (Fig. 6). The inhibitory capacity of crude epiprotein decreased rapidly on heating for 5 minutes above 59° (Fig. 7). The inhibition is observed immediately after mixing OTCase extract with epiprotein (Fig. 8), whatever the amount of epiprotein (Fig. 9). Such a behavior shows that the process is not catalytic, but distinct from an enzymatic transformation, such as the conversion of glycogen phos-

7

ARGININE METABOLISM IN SACCHAROMYCES

0

200 400 600 Arbitrary units of epiprotein

FIG. 6. Increasing amounts of crude epiprotein from cells grown on arginine as only nitrogen nutrient are added to crude ornithine transcarbamylase (OTCase) (from cells grown in ammonium medium) corresponding to 2 mg of proteins

100

g50| u o "o

I

io

i

5h

40

20

60

•c

FIG. 7. Thermodenaturation of epiprotein. A crude epiprotein preparation (8 mg protein ml - 1 ), in Tris 0.1 M, pH 8, is heated for 5 minutes at different temperatures and tested for inhibitory activity as in Fig. 6 {25). OTCase, ornithine transcarbamylase.

EPI

jiol E o

OTC

/

/ ^ OTC ♦ EPI at /=o

V ~3

ö

Γ

6

1

2

t

1

6

I

8

1_ ...

ÏÔ"

Incubation time, minutes at 15°C

FIG. 8. Inhibition of ornithine transcarbamylase (OTCase) by epiprotein is rapid. Crude preparation of OTCase and epiprotein (300 units, see Fig. 6)

8

J. M. WIAME

Incubation time

(minutes)

FIG. 9. Inhibition is not a catalytic process. As in Fig. 8, but increasing amounts of epiprotein are added at 0 time. O—O, no epiprotein; # — # , 40 units; Δ — Δ , 162 units; □ — D , 650 units (25).

phorylase observed in mammalian tissues (9, 12) and of glutamine synthetase in Escherichia coli (18,23,24,47). The inhibited mixture was shown to recover a large part of its activity by dialysis or by a treatment with beef-liver arginase, suggesting that arginine in the crude extract con­ tributes to the inhibition. It should be mentioned that arginine has no effect on a partially purified OTCase alone, but that ornithine, one of the substrates, inhibits the enzyme at concentrations higher than 7 X 10~3 M. (This is the pool of ornithine in cells grown in minimal medium.) OTCase

0.20 0.15

ft \

~~ 0.10 0.05

L·^ 1

02

I

I

.

0.5 1/ornithine ]Q~2M

I

1

1

FIG. 10. Inhibition of ornithine transcarbamylase (OTCase) by ornithine (reciprocal plot). # — # OTCase alone; O—O, with addition of 60 units of epiprotein; Δ — Δ , with 60 units of epiprotein and 10~3 M arginine (25).

9

ARGININE METABOLISM IN SACCHAROMYCES

~>4 u nj Ci)

o 2 hO

10"A

5 x10" 4 Arginine

10"3/tf

FIG. 11. Effect of arginine on the inhibition of ornithine transcarbamylase (OTCase) in presence of 5 X 10~3 M ornithine and 125 units epiprotein {25).

is an enzyme which shows an inhibition by excess of substrate. Addition of a partially purified epiprotein amplifies this effect (Fig. 10). If arginine too is added, there is a strong additional inhibition, which is already detectable at 10"5 M and increases to 5 X IO -4 M (Figs. 10 and 11). Arginine and ornithine are thus acting as effectors in the inhibition; other amino acids have no significant action. A purification of the epi­ protein (180 times) is described in Table II. The purification includes a thermodenaturation which was studied in some detail as a function of arginine concentration (Fig. 12). Arginine is a strong protector, a fact suggesting that this effector acts by binding to the epiprotein. Arginine not only protects, but increases, the

IOOI

50^

I 30 -C c

^

10

20

40

60°C

FIG. 12. Thermodenaturation of epiprotein as a function of arginine concentra­ tion {25).

10

J. M. WIAME TABLE II PURIFICATION OF ;EPIPROTEIN 0 l

Steps 0 1. Crude extract 2. Acid precipitation*

Fractions

a. Supernatant b. Washing of pellet c. Pellet

a. Supernatant 3. Thermodenaturation at 63°C in presence of 10"1 M b. Pellet L-arginine 4. First (NH 4 ) 2 S0 4 fractionation« 5. 2nd fractionation by precipitation of fraction 4b with (NH 4 ) 2 S0 4

a. 55-90% b. 30-55% c. 0-30% a. b. c. d. e. f.

30-35% 35-40% 40-45% 45-50% 50-55% 55-60%

Volume (ml)

Total Epiproteic activity 6 protein Specific Total (mg) 2500 890

95 214

—

239,000 188,000 21,900

25

975

18,200

19

43 25

250 612

269,000 18,500

1,070 30

50 42 7.7

6,850 187,000

—

137 4,450 292

6.8 6.4 8.2 6.6 9.5 2.2

2,400 16,700 145,000 24,000 687 138

850 2,330 17,500 3,640 72 61

24.5 33 6.5

24 6.6 4.3 1.01 1.03 1.02 1.0 1.04 0.98

—

a

Data from Messenguy {25). The purest fraction, 5c, corresponds to a 180-fold purification and a yield of 54%. The quantity present in strain 1705 when grown in L-arginine as the sole source of nitrogen is 100 arbitrary units. The activity is measured by inhibition of ornithine transcarbamylase in presence of 5 X 10~2 M L-ornithine and 10"3 M L-arginine in a range of inhibition smaller than 50%; in this range, the inhibition is almost a linear function of epiprotein concentration. 0 All steps except step 3 in presence of L-arginine 5 X 10~3 M. d Acidification to pH 5 with acetic acid; the supernatant is neutralized with Tris, free base. e By solubilization. b

epiproteic activity in step 3 of purification (see Table II). Ornithine is of little help in protection, and most probably acts only through OTCase (see also Section II, D). C. The Nature of the Epiprotein and Its Binding with Ornithine Transcarbamylase (25, 28)

The participation of ornithine as an effector calls attention to the meta­ bolic peculiarity of this intermediate : it appears as an intermediate in the biosynthesis as well as in the degradation of arginine. For this reason,

11

ARGININE METABOLISM IN SACCHAROMYCES

5x10-2 arginine

FIG. 13. Critical temperature of epiproteic activity ( # — # ) and arginase activity (O—O) {25). See Fig. 12 for the operational definition of critical temperature.

the possibility that an enzyme of catabolism could be identical with the epiprotein was investigated. It was observed that during the purification of the epiprotein, the purification of arginase runs in parallel. Especially striking was the fact that the thermostability as a function of arginine concentration gave identical critical temperatures (Fig. 13). The ornithine transaminase activity did not behave similarly, and the pyrroline carboxylate dehydrogenase could not be involved because it is constitutive (89). Finally, the molecular weight of the material which carries the epiproteic activity is identical to that of the arginase activity (114,000) when esti­ mated by molecular sieving (Fig. 14).

o

-3000

c

-2000

ï il Λ

ce.E H-

û>

° o -1000

C Q. ZD ft)

1

10

;\

_

Λ

jL ïï\k 30 fractions

*

Λ 50

FIG. 14. Elution of arginase and epiprotein on Sephadex G-200 superfine with Tris 0.05 M, KC1 0.2 M, and arginine 5 X IO -3 M. The peaks of the two activities, arginase (O—O) and epiprotein (▼—T), are in the same fractions, after the marker, lactic dehydrogenase ( □ — □ ) (mol. wt. 135,000), and before alkaline phosphatase (V—V) (mol. wt. 80,000). 1 1— blue dextrane. — Δ—Δ— cytochrome c. The molecular weight of arginase is about 114,000

12

J. M. WIAME

800l·

o400

fractions

FIG. 15. Elution of a mixture of arginase (O—O) and ornithine transcarbamylase ( # — φ ) as in Fig. 14, but no arginine. The molecular weight of ornithine transcarbamylase (OTCase) is about 120,000. See also Fig. 14 for symbols (28).

Taken together, these data suggest at least a strong relationship, if not identity, between the two proteins. The study of arginaseless mutants gave further information. One of these mutants (AGI) retains an epiproteic activity, whereas two others (AG2 and AG3) were not active. Except for the possibility that the genes for arginase and epiprotein are contiguous and mutation in one affects the other by polarity, it follows that epiprotein is identical with arginase, and that the AGI mutant contains a mutated protein in which the catalytic function has been lost, while the regulatory function is retained. A definite proof of identity of the molecules which bear regulatory and arginase activities is given by the strong binding between arginase and

^600 8 400 y 200 o

-

aA

>

+ ?,

-

\ .'I\ ^

-1

III

10

fl

y

;M\

QT

CD

200 < 1 È

fractions

l\ --moo s 1 \- c

;

1

40

FIG. 16. Elution of a mixture of arginase (O—O) and ornithine trans carbamylase (OTCase) ( · — # ) . Same as Fig. 15, but in presence of ornithine 1.5 X IO"2 M and arginine IO-3 M. A common peak of the two activities appears before the lactic dehydrogenase ( □ — □ ) , corresponding to molecular weight between 180,000 and 200,000. An excess of arginase remains at its usual molecular weight (110,000). See also Fig. 14 for symbols {28).

13

ARGININE METABOLISM IN SACCHAROMYCES

-h 200

ouur

1

Έ.

f, 400

H 800 tj

À 200

-Uoo g

H

A * 40

fractions

J

Û)

O)

1 ^

60

FIG. 17. Elution of a mixture of arginase (O—O) and ornithine transcarbamylase ( · — # ) . Same as Fig. 15, but arginine 10~3 M and ornithine 2 X 10~3 M. See also Fig. 14 for symbols [Gevers and Messenguy, cited in {25) ].

OTCase when both ornithine and arginine are present, and the absence of this binding when either ornithine or arginine are absent. On Sephadex G-200, OTCase is eluted in fractions corresponding to a molecular weight of 120,000 and arginase, 114,000 (Fig. 15). In the presence of 1.5 X 10~2 M ornithine and 10~3 M arginine, arginase is eluted in two peaks. One of these peaks corresponds to a molecular weight around 180,000 and also contains all the OTCase. The second peak corresponds to an excess of arginase, free of OTCase, in its usual elution fraction (Fig. 16). In the presence of 10~3 M arginine, ornithine begins to provoke binding at 2 X 10"3 M (Fig. 17). With 1.5 X 10"2 M ornithine, a partial binding occurs with 2 X 10~5 M arginine. This shows that arginase of S. cerevisiae has regulatory properties and, together with ornithine transcarbamylase, forms an allosteric system with a substrate of each of the enzymes as effectors. The second substrate of OTCase, carbamyl phosphate, does not seem to be involved in this regulation. It reactivates an inhibited OTCase, but at high and unphysiological concentrations. D. Physiological Significance {28) Identification of the regulatory protein as arginase allows us to interpret the meaning of this regulation. The inhibition of OTCase by arginase avoids the establishment of a meaningless urea cycle which costs 5 energy-rich bonds (see Fig. 1). When arginine is added to minimal medium, or when cells are shifted from minimal medium to a medium in which arginine is the nitrogen nutrient, the synthesis of OTCase is much reduced and OTCase is progressively diluted out by growth. In the meantime, ornithine could be recycled into arginine, either with the carbamyl phosphate destined to the pyrimidine synthesis or with the carbamyl phosphate for arginine, since for the latter no feedback inhibition was detected.

14

J. M. WIAME

It must be mentioned that, in the state of repression, the level of OTCase is by no means negligible when compared to other biosynthetic enzymes {45), and one can see in this inhibited residual OTCase a potential capacity for arginine biosynthesis when the effectors are depleted. By such a mech­ anism, one could avoid a long lag phase and even absence of growth, such as that described in Proteus, when this organism is transferred from condi­ tions of repression of sulfur amino acid metabolism to minimal medium {6). This may be the result of the well known strong degradative capacity of Proteus for sulfur amino acids which cannot be compensated by a minimum of residual syntheses. There is no reciprocal effect of OTCase on arginase activity. Arginase does not affect other enzymes of the biosynthetic pathway. E. Specificity of Binding {25, 26) The study of cross reactions between heterologous arginases and OTCases shows that the specificity of binding is a function of both proteins. The occurrence of suppression of OTCase activity in situ (described in Section II, A) in all species of the genus Saccharomyces suggests that all these OTCases are sensitive to the respective homologous arginases. The species tested were S. cerevisiae, S. pastorianus, S. italicus, S. bayanus, S. fragilis, S. turbidans, S. oviformis, S. dobzanskii, S. chevalieri, and S. diastaticus. Arginases from S. chevalieri and S. italicus inhibit the OTCase from S. cerevisiae in cell-free systems (Section II, B), and OTCases from S. carlsbergensis, S. fragilis, and S. italicus are sensitive to the arginase of S. cerevisiae. One may suppose that cross reactions between heterologous arginases and OTCases are general in the genus Saccharomyces. No in situ suppression occurs in yeasts like Candida utilis, Pichia fer-

Pichia fermentans

40

^> fc|20

V)

σ o 1-

; l ί H V* ί '· li «,ί 20

♦ 1 * ;.

60 fractions

FIG. 19. Elution of a mixture of ornithine transcarbamylase (OTCase) from Debaryomyces globosus (mol. wt. 88,000) in presence of arginase from Saccharomyces cerevisiae (mol. wt. 114,000), in presence of ornithine 1.5 X 10~2 M and arginine IO-3 M. Binding is shown in the common peak for the two activities, corresponding to a molecular weight of 160,000. See also Fig. 14 for symbols {25).

mentons, and Debaryomyces globosus. The synthesis of OTCase is not repressible on the first of these yeasts, but it is in the two others. In cell-free extracts, the arginase from S. cerevisiae does not inhibit most OTCases. These include the bacterial OTCases from Escherichia coli and Pseudomonasfluorescens (the anabolic enzyme) and the yeasts OTCases from Candida tropicalis, C. utilis, and Pichia fermentons. The OTCases from C. utilis and P. fermentons have hyperbolic saturation functions, but the OTCase from D. globosus shows inhibition by excess of ornithine (Fig. 18). The first two OTCases are insensitive to the arginase of Saccharomyces, but that of D. globosus is sensitive, although this species does not show suppression and, accordingly, is insensitive to its own arginase. D. globosus OTCase binds with S. cerevisiae arginase. The molecular weight of D. globosus OTCase is 85,000, and in the presence of Saccharomyces arginase it moves with it to a common peak which is eluted in a fraction corresponding to molecular weight 160,000 (Fig. 19). There is no such indication of binding between Pichia fermentons OTCase and the arginase of Saccharomyces. Again, the capacity for an OTCase to be inhibited by excess substrate appears to be required for inhibition by arginase. We propose that an enzyme able to impose its action on another by binding be called an epienzyme: epiarginase in the present case. F. The Regulatory Site of Ornithine Transcarbamylase {28a, 32a, J^S) 1. DISTINCTION OF T W O SITES

The saturation curve of OTCase with ornithine begins as a hyperbolic function, followed by inhibition. This inhibition can be determined quanti­ tatively by comparing experimental activities with extrapolated hyperbola

16

J. M. WIAME

20

40 60 Ornithine x IO"3 M

FIG. 20. Ornithine transcarbamylase (OTCase) activity as a function of ornithine (O—O). Inhibition of OTCase (% I) by ornithine ( · — · ) calculated from the extrapolated hyperbola ( X — X ) . Same in presence of putrescine 5 X 10~2 M (H h) (28a).

in a Lineweaver and Burk plot (Fig. 20). This inhibition appears co­ operative. Substances that may simulate ornithine could be useful for analysis of both the catalytic and inhibitory functions of ornithine. It has already been mentioned that epiarginase, or epiarginase plus arginine, seems to act by an amplification of the property of ornithine to inhibit OTCase activity at high concentrations (see Fig. 10). A more accurate study of such a relation must avoid the slight conversion of arginine into ornithine. This can be done by using an epiarginase from the AGI mutant: this protein has a regulatory, but no catalytic, activity (see Section II, B).

epiarginase* 10~3/if arginine /♦epiargif

0 0.1

0.5 , 1/ornithme IO"3 M

FIG. 21. Inhibition of ornithine transcarbamylase (OTCase) by an epiarginase from mutant AGI which has no arginase activity. This allows the study of the apparent affinity of OTCase at low concentration of ornithine in presence of epiarginase and ornithine (to be compared with Fig. 10) (28).

17

ARGININE METABOLISM IN SACCHAROMYCES

FIG. 22. Competitive inhibition of ornithine transcarbamylase a, 7-diaminobutyrate (28a).

(OTCase)

by

The results are reported in Fig. 21 (to be compared to Fig. 10) ; obviously epiarginase or epiarginase plus arginine does not affect the apparent affinity, but only the inhibition by excess of substrate. From this, one may predict distinct catalytic and inhibitory sites for ornithine. a,7-Diaminobutyrate is known to be a competitive inhibitor of OTCase from Neurospora crassa (17). This is also true for the OTCase from S. cerevisiae, and simple competitiveness occurs without additional inhibition at high concentrations of the analog (Fig. 22). Putrescine, the ornithine analog lacking the carboxylic group, decreases the inhibition by excess of substrate and, contrary to a, 7-diaminobutyrate, is not a competitive inhibitor of OTCase activity (Fig. 20). At a low concentration of ornithine, putrescine is without activity. Most probably putrescine binds to the regulatory site, but cannot cause the inhibition. Hydrogen ion is a noncompetitive inhibitor of OTCase; it does not I

0

PH7

0.5 1.0 1/ornithine IO - 3 M

2.0

FIG. 23. Noncompetitive inhibition of ornithine transcarbamylase by hydrogen ion. The inhibition by ornithine disappears when hydrogen ion concentration increases {28a).

18

J. M. WIAME

affect the apparent affinity for ornithine, but strongly affects inhibition by excess of ornithine (Fig. 23). 2. DESENSITIZATION

Desensitization has been observed in many allosteric enzymes and was an early indication for a distinction between regulatory and catalytic sites, as well as for the hypothesis that regulatory interactions are indirect (7, 13, 22, 30). Inhibition of yeast phosphofructokinase by excess of ATP has also been proved to be exerted at a site which is distinct from the catalytic one by the elegant work of Salas, Salas, and Sols (35), showing that inhibition is abolished by a limited trypsin digestion. a. Acetylation ofOTCase. The catalytic site of beef-liver OTCase probably contains a tyrosine residue which can be acetylated by acetylimidazole (15). With the thought that recognition of ornithine at two distinct sites might use similar amino acid residues, acetylation of the yeast enzyme was per­ formed in the presence of ornithine, which should preferentially protect the catalytic site which has a higher affinity for ornithine than the regulatory site. After this treatment, 80% of the catalytic activity was retained while the inhibition by excess of substrate disappeared. The affinity for ornithine was unchanged (Fig. 24) and both inhibition by arginase (Fig. 25) and binding tested by molecular sieving (Section II, C) were lost. ò. Heat. Heating OTCase under conditions described in Fig. 26 provokes a decrease in inhibition by excess of ornithine, and the function of saturation

acetylated

0 1 , 2 1/ornithme 10" J M

FIG. 24. Desensitization of ornithine transcarbamylase by acetylation. Treatment by iV-acetylimidazole 2 X 10 -1 M for 1 hour in presence of 50 X IO-3 M ornithine in triethanolamine 0.02 M, pH 8 (28a, 45).

19

ARGININE METABOLISM IN SACCHAROMYCES

o

100

^

Ci

r.

acetylated OTCase

2 50 native OTCase

5 0

1

200

,

1

400 Arginase

_. _J

600

FIG. 25. Action of epiarginase on ornithine transcarbamylase (OTCase) in presence of 15 X IO -3 M ornithine and IO -3 M arginine (28a).

20 40 ornithine x 10 3 M

FIG. 26. Partial desensitization by heating ornithine transcarbamylase in presence of ornithine 5 X 10"2 Af, at 58°C in Tris 0.1 M, pH 6.5 for various times. Activity measured at pH 8 and 15° {28a).

0.5 1.0 1/ornithine IO"3 M

15

FIG. 27. Data of Fig. 26 in reciprocal plot. The-apparent affinity is unchanged, and the increased activity is due to the loss of inhibition (28a).

20

J. M. WIAME

1.5h

^

1.0h

^

j y ^

0.5^

0

0.5 1.0 1.5 1/ornithine 10 -3 M

FIG. 28. Ornithine activation rather than inhibition with ornithine transcarbamylase (OTCase) from revertant strain RF90 (dialyzed crude OTCase) {28a).

approaches a hyperbolic form (Fig. 27). The catalytic function (Fmax extrapolated from low ornithine concentration) is not affected. c. Mutation. Among spontaneous revertants of the OTCase" mutant MG802, the revertant RF90 does not show inhibition by excess of ornithine. On the contrary, it shows activation by excess of ornithine (Fig. 28) ; this curious behavior can be compared to that of a mutant at prephenate dehydratase of Bacillus suhtilis which, contrary to the wild-type enzyme is activated by phenylalanine (8). The sensitivity of OTCase toward arginase is much reduced in the RF90 mutant (Fig. 29).

0

J

200

I

L

ωθ 600 Arginase

FIG. 29. Reduction of inhibition of crude ornithine transcarbamylase (OTCase) from strain RF90 compared to OTCase from wild type. Test as usual: 1.5 X 10~2 M ornithine, 10~3 M arginine, and various amounts of arginase {28a).

21

ARGININE METABOLISM IN SACCHAROMYCES

III. Common Genetic Elements in the Control of Biosynthetic and Degradative Enzyme Synthesis* A. Reciprocal Balance between Biosynthetic Enzyme Syntheses (3, 34, 39, 40, 45)

and

Degradative

1. W I L D - T Y P E STRAIN

A number of compounds added to minimal medium repress the synthesis of ornithine transcarbamylase. These are L-arginine, DL-homoarginine, L-ornithine, a, γ-DL-diaminobutyrate, and L-lysine. Growth on glutamate as the nitrogen nutrient also causes repression. All these conditions, without exception, induce the synthesis of arginase. The parallelism is striking and suggests a common regulatory element (Table I I I ) . This element does not appear to be a unique metabolite like arginine, because ornithine at high concentration (ammonium plus 1000 Mg ml" 1 ornithine), while fully active in repression or induction, reduces the pool of arginine from 18 X 10~3 M to 5 X IO -3 M. Nor is it ornithine, since repression as well as induction occurs with arginine (ammonium plus arginine), together with a reduction of the ornithine pool. The fact that arginine induces ornithine transaminase in arginine-less strains also shows that, even in this case, arginine does not act through ornithine. Furthermore, the active analogs can hardly be transformed either into arginine or into ornithine. Hence, arginine and ornithine must have separate actions, eventually at distinct sites, through a common regulatory element. 2. argR

MUTANTS

Three unlinked mutations with similar phenotypes, argRI, ar^RII and ar^RIII, have been selected on the basis of resistance to a small amount of canavanine (8 Mg ml -1 ) in the presence of ornithine (200 Mg ml - 1 ). Under these conditions, ornithine represses OTCase in the wild-type strain, and * Nomenclature: The nomenclature proposed by Demerec et al. (11) for bacterial genetics has been adopted. argB, argC . . . correspond to mutations in enzymatic steps in arginine biosynthesis; cargA and cargB correspond to mutations in the first (arginase) and the second (ornithine transaminase) steps of arginine catabolism, respectively. The wild-type alleles are designated argrB+, or simply + if no confusion is possible. When mutations concern an unknown biochemical process, or a regulatory mechanism, they are designated by a capital letter from the second half of the alphabet, as in argR. If different classes of mutations have obvious similarities, a roman number distinguishes them, as in argKl, argKll, and argKHI. Individual mutations are designated by arabic ciphers (3). Operator mutation for a gene as cargA in its wild-type state cargA+ is designated cargA+0 {20). If the ultimate product of a gene is not an enzyme, it is desig­ nated by the name of the gene in capitals; thus, A R G R I + is the product of argKI+.

S1278b

Strain

Wild t y p e

Genotype 6

1 2 3 4 5 6 7 8 9 10 11 12

Expt. no.

N H 4 + + DL-homoarginine N H 4 + + DL-a, 7-diaminobutyrate L-Ornithine L-Arginine L-Glutamate L-Proline Urea Chemostat, NH 4 + IO" 3 M

NH 4 + + L-lysine

NH 4 + N H 4 + + L-arginine N H 4 + + L-ornithine

Nitrogen nutrients available

200 200 600

300 150

110

Generation time (minutes at 29°C)

6 to 8 18 21 22* 19* 21 110 200 40 15 19 32 to 40

Arginase

7.5

0.03 0.24

L-Ornithine-δtransaminase

28

29 to 32 2 12 9 15 15 12 1 19 37

OTCase

Enzyme activities (jumoles product formed, mg protein - 1 , hour" 1 , at 30°C)

CATABOLIC ENZYMES IN W I L D - T Y P E AND argR M U T A N T *

R E P R E S S I O N O F O R N I T H I N E TRANSCARBAMYLASE ( O T C A S E ) AND INDUCTION O F

TABLE III

H

h-1

3 >

g

ÇH

to

LO

{argRÏ2 ] largrRIIlOJ

13 14 15 16 17 18 19 20 21 22

NH 4 + NH 4 + + L-arginine NH 4 + + L-ornithine NH 4 + + L-lysine NH 4 + + DL-homoarginine NH 4 + -f DL-a, 7-diaminobutyrate L-Arginine or L-ornithine L-Glutamate Urea Chemostat, NH 4 + 10~3 M 200 600

1000

° Data from (34, 39, 45) and unpublished. 6 See footnote p. 21. c Induction first observed by Bourgeois and Thouvenot (6). d Induction first observed by Whitney and Magasanik (personal communication).

7000a

15

15

3 3 3.5 3.5 4 4

0.026 0.032

43

39 to 44 37 42 39 41 45

to

GO

O

O

>

o o M

>

g

5

W Ο

3H

2 S

>

24

J. M. WIAME

the pool of arginine decreases from 20 X 10~3 M to 12 X IO -3 M. In the mutants, repression is impaired, the pool of arginine on minimal medium is 25 X 10~3 M, and ornithine raises the arginine pool to 37 X IO -3 M, which competes with canavanine (3, 34). The argR mutations provoke derepression and constitutivity of OTCase and some other enzymes in the pathway (Table I I I ) . These mutations are largely recessive in diploids. However, in tetraploids with only one dose of wild-type allele and three doses of argR, the dilution of the wild-type product by a factor of four reduces repressibility (17b). argR mutations have been interpreted as regulatory mutations affecting the synthesis of a heteropolypeptidic aporepressor acting on the synthesis of five enzymes of the biosynthesis out of a total of nine. Unexpectedly, these mutants are unable to use arginine, ornithine, or citrulline as a sole source of nitrogen because the catabolic enzymes arginase and ornithine transaminase are induced by none of the effectors already men­ tioned, including growth on glutamate (40). Hence, argR mutations appear to affect the common element mentioned above (Section III, A, 1), and their action is not restricted to the control of biosynthetic enzyme synthesis. The requirement for argR+ products to induce arginase and ornithine transaminase is typical behavior of positive control (36). However, it is not expected that one macromolecule could act negatively on an anabolic gene and positively on a catabolic one. This prompted us to look for additional regulatory elements for catabolism with the idea that a positive control could be due to a sequence of two negative controls. B. Negative Type of Control of Degradative Enzymes (17a, 45) 1. HYPOTHESIS OF TWO SUCCESSIVE NEGATIVE CONTROLS

An apparent positive control of arginine catabolism by argR+ products which might result from the sequence of two negative steps is represented in Fig. 30A-C. This hypothesis predicts classical regulatory elements, i.e., repressor and operators, for arginase and ornithine transaminase. Since the structural genes for these two enzymes, cargA and cargB, are not linked, two distinct operators would be necessary. The second aspect of the hypothesis con­ cerns the transmission of the signal. The argR+ product transmits metabolic signals to the anabolic genes in the usual way, the argR+ product being activated by arginine or ornithine. On the other hand, the metabolic signal also reaches the catabolic regulatory circuit through argR+ products which, when activated, either inhibit the activity of the gene coding for the

A R G I N I N E METABOLISM I N

SACCHAROMYCES

25

repressor of catabolism (Fig. 30B) or, as would an inducer, deactivates the repressor of cat abolie enzymes (Fig. 30C). 2. CONSTITUTIVE MUTANTS FOR ORNITHINE TRANSAMINASE AND ARGINASE

a. Method. The hypothesis presented in Fig. 30 can be analyzed by a search for constitutive mutants for catabolic enzymes which overcome the absence of induction resulting from arg~R mutations. Reversion of argR to wild-type allele can be avoided by using a double mutant an/RI, argRII (strain 7000a) as mother strain for the selection. The recovery of normal growth on ornithine as the sole source of nitrogen requires constitutivity only of ornithine transaminase, since the third enzyme of the pathway, pyrroline carboxylate dehydrogenase is already constitutive (39). The recovery of normal growth on arginine requires the constitutivity of both ornithine transaminase and arginase. Dominance or recessiveness expected for operator and for repressor mutations, respectively, can be analyzed directly in diploids homozygotic for argRl and ar^RII, resulting from a cross with the strain 120c which is identical with the parent strain of the mutants, 7000a, except for the mating type. Once selected, each new mutation is transferred into a wild-type genome by crosses with a wild-type strain. Selection of mutants growing normally on ornithine gave two classes of mutations when tested for dominance. A mutant like DH43 bears a domi­ nant mutation. This mutation was transferred into a wild-type genome in the segregant 7051a. This mutation is analyzed in Section III, B, 2, b. Mutants like DH29 and DH51 bear recessive mutations present in a wildtype genome in segregants 7010d and 7142b, respectively. These mutations are analyzed in Section III, B, 2, d. Attempts to select mutants growing normally on arginine fail, unless the parent already bears a constitutive mutation for ornithine trans­ aminase, as is the case for strain DH43. Two new mutants growing normally on arginine, DH4314 and DH437, have been analyzed. Each of the two mutations is present in a wild-type genome in segregants 7074a and 7093a, respectively. These mutations are described in Sections III, B, 2, c and e. b. Operator for Ornithine Transaminase. The dominant mutation present in mutant DH43 and segregant 7051a has the properties of an operator mutation for ornithine transaminase (mutation cargB+0) : the enzyme level is some 100 times as high as in the wild-type strain, when both strains are grown on minimal medium, and the mutation is without influence on the level of arginase (Table IV, Expt. 3a, 3b, 3c; cf. Expt. la, lb, lc).

26

J. M. WIAME

ornithine transcarbamylase inactive ARGR*

arg F*

(

\

active CARGR+—\

arg RUT*

-|——i 1_ 0 + carg A*

cargR* H

FIG.

1 y 0 + cargB +

30A

1

μ_

0 + carg A + cargR*

ornithine transaminase

,?

FIG.

30B

0 + carg B+

The heterozygous diploid has half the level of ornithine transaminase, corresponding to a dilution by a factor of two, as the wild-type genome does not contribute to a detectable extent under these conditions (Table IV, Expt. 5a; cf. Expt. 3a). The proximity between mutation cargB+0 and the structural gene for ornithine transaminase, carg'B has been analyzed, and a strong linkage is apparent. A crossing-over between the constitutive-operator mutation

27

ARGININE METABOLISM IN SACCHAHOMYCES

arginine 9r ornithine inactiv e + ARGR:

I -^

active * ARGR +

inactive CARGR

•i

arg F*

arginase

active CARGR+

arg RIE*

j

Î 1

0 + cargA*

v

ornithine transaminase cargR*

,,, ί,, 0 + carg B +

FIG.

30 C

FIG. 30. Hypothetic scheme for an apparent positive control resulting from a sequence of two negative steps, H—ι—H-, gene and operator; - ^ w > ARGR, ultimate o+ product of a gene written in capitals, unless it is an enzyme; argF+, a structural active gene in the anabolic pathway; cargA+, an active gene in catabolic pathway. Roman numerals I, II, III indicate distinct genetic classes, each indispensable but with very similar functions. They are supposed to form a heteropolypeptidic aporepressor ARGR. (A) In the absence of arginine or ornithine. (B) In the presence of arginine or ornithine. Hypothesis of inhibition of production of the aporepressor for catabolism, cARGR. (C) As in scheme B, hypothesis of an inhibition of the activity of the aporepressor.

adjacent to the active transaminase cargB+0 gene and a transaminasedefective locus with repressible operator cargBO+ should give rise to a segregant cargB+0+ with very low level of transaminase in minimal medium. This analysis did not show such recombination among 16 tetrads analyzed by enzyme determination: only parental types were detected. To perform an analysis on a larger scale, the cross was made in the presence of an argR background obtained by crossing cargB+0, argBlI with cargBO+, argBII. In this case, a recombinant cargB+0+, argRII can be detected by its incapacity to grow on ornithine as nitrogen source. By this method, 104 tetrads were analyzed, giving 97 parental ditypes and 7 tetrads con­ taining 3 segregante unable to grow on ornithine instead of 2. Six of these 7 tetrads contain 3 cargB instead of 2; i.e., they have undergone not a crossing-over, but a gene conversion. If further analysis of tetrads provides a cargB~0~ recombinant, it will be possible to test whether, as expected, there is absence of dominance in the diploid car^B"O~x car^B + 0 + .

,

+

(cargB+0)

)

5a

4a b c d

cargB+0 | argRI2 ^RIIIO,

DH43

Diploid 7051a X 3962c

3a b c

cargB+0

c

7051a

Operator mutations

2a

1 argRI2 argRlIlO 1

7000a

b

la b c d e

Expt. no.

Wild type

Genotype

21278b

Strain

TABLE IV

0.026 0.032

NH4+ N H 4 + + L-arginine N H 4 + + L-ornithine L-Arginine or L-ornithine

1.3

2.8 2.9 3.1 4.0

NH4+ N H 4 + + L-arginine N H 4 + + L-ornithine L-Ornithine NH 4 +

3.2 3.4 3.3

NH4+ N H 4 + + L-arginine N H 4 + + L-ornithine

no growth

0.03 0.24 0.2 7.5 2

L-Ornithine-5transaminase

NH4+ N H 4 + + L-arginine N H 4 + + L-ornithine L-Arginine L-Ornithine

Nitrogen nutrients available

3.3 4.2 4.2 20

—

9

3 3.5 3

6-8 18 21 200 110

Arginase

Enzyme activities (μηιοΐββ product formed, mg protein -1 , hour - 1 at 30°C)

CONSTITUTIVE MUTATIONS FOR ORNITHINE TRANSAMINASE AND ARGINASE

3 >

g

JH

00

29

ARGININE METABOLISM IN SACCHAROMYCES

O IO O O ^H o * "M i-t CO

O àO

TH

H

co co O

Ì4

c3

cu

+ 'Sb.'ä

?

.'§;§

•3

+ tì -g

o3 ^D CO

2 - tì tì tì

tì

Ci (M

ffl Û

X5 (M

iO

Q

6

α

See footnote page 32. Same genotype as 7010d.

Diploid 7093a X 21278b

7093a

Catabolito repressor for arginase?

Diploid 7010d X S1278b

Diploid 7142b X 7143dft

Strain

, +

(cargAli i

cargrAR

{ + J

cargRl

cargRlj

cargR2

Genotype

16a

15a b e

Nitrogen nutrients available

NH 4 +

NH4NH=>), catabolism (^ ■>-), and action of effectors ^ ) which contribute to ornithine transcarbamylase (OTCase) inhibition (28).

ORNITHINE

TRANSCARBAMYLASE Ornithine

Regulatory,—.

3

3x10~ /W

Carbamyl phosphato

Λ

20 χ1θ" Λ/

site-^ /buried /binding CR-^l Orn-Qi

v^/area ·/

Ä_ L

__

ARGINASE

b

[Arginine site

FIG. 32. Attempt to describe epiarginase and ornithine transcarbamylase inter­ action (28a).

36

J. M. WIAME

nature of the two binding areas, and almost nothing is known about the site for arginine except that it is located on the arginase molecule and does not need the catalytic process to be intact. It is not known whether the regulatory site and the site of binding of arginine to initiate catalysis are distinct. The contribution of the ornithine site begins to emerge, thanks to the observation of two phases in the activity of OTCase as a function of ornithine concentration. A classical hyperbolic function of saturation is followed by a phase of inhibition with a cooperative effect. The specificity of this inhibition is so great that it can hardly be explained by the trivial overcrowding of the catalytic site. This inhibition occurs with OTCase from Saccharomyces and from Debaryomyces globosus, which are sensitive to epiarginase. It does not occur with OTCases insensitive to epiarginase. Indeed, the capacity to be inhibited by excess of ornithine appears to be required for an OTCase to be sensitive to epiarginase. Acetylation in the presence of ornithine and mutations lead to OTCases which have simul­ taneously lost inhibition by excess of ornithine and sensitivity to epi­ arginase. The regulatory site appears distinct from the catalytic one. One of the most direct evidences for this distinction is that partial heat desensitization may be obtained without change of the catalytic capacity. Fig. 32 represents the main steps in OTCase-epiarginase interaction. C. Other Considerations A common protein involved in induction of catabolic enzymes and in repression of OTCase is suggested by the strict identity of the effectors of both regulations and by argil mutations which simultaneously abolish repression of OTCase and synthesis of inducible enzymes. arv/R mutations belong to three genetic loci, argBl, argKII, ar^RIII, which appear to con­ tribute to the synthesis of a heteropolypeptidic aporepressor, ARGR, which might be the common regulator for induction and repression. This ambiv­ alent function might be exerted by a direct classical negative control on the synthesis of OTCase and by a sequence of two negative controls of the catabolic enzymes, as presented in Fig. 30. The existence of operators for arginase and ornithine transaminase, and of a pleiotropic aporepressor (CARGR) for the two enzymes (which is expected on the basis of this hypothesis) is reported in Section III, B, 2, b. Furthermore, this hypothesis foresees that the signal for induction, the first part of the mechanism, is transmitted by the common ambivalent repressor ARGR. Data reported in Section III, B, 2, d agree with this view: argil mutations abolish induc­ tion. Hence, it seems that the apparent positive control of the ambivalent element on the catabolic enzymes (which was difficult to reconcile with its negative control on biosynthetic enzymes) is obtained by the combination

ARGININE METABOLISM IN SACCHAROMYCES

37

of two negative actions: the inhibition of an inhibitor. One may wonder whether other cases of positive control could not be of this sequential doubly negative type. As presented here, the ambivalent repressor is a tool for a mechanism of balanced exclusion of enzyme syntheses involved in concurrent pathways. Induction of catabolic enzymes is far from being sufficient for the use of arginine or ornithine as sole source of nitrogen. This is true also for catabolic derepression (see Table I). Both induction and catabolite derepression are necessary together. However, in the presence of operator mutations, a high level of constitutivity compatible with growth on arginine or ornithine is obtained. This suggests that these operators may be implicated in induction and catabolite repression. Indeed, a mutation which is largely recessive and specific for arginase brings the level of arginase to the constitutive-operator level, and may open new possibilities in the study of nitrogen catabolite repression of eukaryotes. ACKNOWLEDGMENTS

I thank Academic Press for permission to reproduce Figs. 2-5 from Biochemical and Biophysical Research Communications (4) and Figs. 20, 22, and 24 from the Reports to the 10th Congress of Microbiology, Mexico, 1970, in press {45) y as well as the editors of FEBS Letters for Figs. 14, 15, 16, and 31 and European Journal of Biochemistry for Figs. 20, 22, 23, 24, 25, 26, 27, 28, 29, 32. REFERENCES

1. Béchet, J., Grenson, M., and Wiame, J. M., Arch. Int. Physiol. Biochem. 70, 564 (1962). 2. Béchet, J., Grenson, M., and Wiame, J. M., Arch. Int. Physiol. Biochem. 73, 137 (1964). 3. Béchet, J., Grenson, M., and Wiame, J. M., Eur. J. Biochem. 12, 31 (1970). 4. Béchet, J., and Wiame, J. M., Biochem. Biophys. Res. Commun. 21, 226 (1965). 5. Bourgeois, C. M., and Thouvenot, D . R., Eur. J. Biochem. 15, 140 (1970). 6. Bourgeois, S., Wiame, J. M., and Lelouchier-Dagnelie, H., Biochim. Biophys. Acta 38, 136 (1960). 7. Changeux, J. P., Cold Spring Harbor Symp. Quant. Biol. 26, 313 (1961). 8. Coats, J. H., and Nester, E. W., J. Biol. Chem. 242, 4948 (1967). 9. Cori, G. T., and Green, A. A., J. Biol. Chem. 151, 31 (1943). 10. De Deken, R., Biochem. Biophys. Res. Commun. 18, 462 (1962). 11. Demerec, M., Adelberg, E. A., Clark, A. J., and Hartmann, P. E., Genetics 54, 61 (1966). lia. Dubois, E., Hiernaux, D., and Wiame, J. M., unpublished. 12. Fisher, E. H., and Krebs, E. G., / . Biol. Chem. 216, 121 (1955). 13. Gerhart, J. C , and Pardee, A. B., J. Biol. Chem. 237, 891 (1962). 14. Gilbert, W., and Müller-Hill, B., Proc. Nat. Acad. Sci. U.S. 56, 1891 (1966). 15. Grillo, M. A., and Coche, M., Italian J. Biochem. 2 8 , 133 (1969). 16. Halleux, P., S talon, V., Piérard, A., and Wiame, J. M., Arch. Int. Physiol. Biochem. 78, 166 (1969).

38

J. M. WIAME

17. Hermann, R. L., Lou, H. F., and White, C. W., Biochim. Biophys. Ada 121, 79 (1966). 17a. Hiernaux, D., Dubois, E., Grenson, M., and Wiame, J. M., unpublished. 17b. Hilger, F., Grenson, M., and Wiame, J. M., in preparation. 18. Hölzer, H., Advan. Enzymol. 32, 297 (1969). 19. Jacob, F., and Monod, J., J. Mol. Biol. 5, 318 (1961). 20. Jacob, F., and Wollman, E. L., "Sexuality and the Genetics of Bacteria." Academic Press, New York, 1961. 21. Mandelstam, J., Biochem. J. 82, 489 (1962). 22. Martin, R. G., / . Biol Chem. 237, 257 (1962). 23. Mecke, D., and Holzer, H., Biochim. Biophys. Acta 122, 341 (1966). 24. Mecke, D., Wulff, K., Lies, K., and Holzer, H., Biochem. Biophys. Res. Commun. 24, 452 (1966). 25. Messenguy, F., Thesis, University of Brussels (1969). 26. Messenguy, F., Béchet, J., and Wiame, J. M., Arch. Int. Physiol. Biochem. 75, 889 (1967). 27. Messenguy, F., and Wiame, J. M., Arch. Int. Physiol. Biochem. 77, 165 (1968). 28. Messenguy, F., and Wiame, J. M., FEBS Lett. 3, 47 (1969). 28a. Messenguy, F., Penninckx, M., and Wiame, J. M., Eur. J. Biochem., in press. 29. Middelhoven, W. J., Biochim. Biophys. Ada 156, 440 (1967). SO. Monod, J., Changeux, J. P., and Jacob, F., J. Mol. Biol. 6, 306 (1963). 31. Monod, J., Wyman, J., and Changeux, J. P., / . Mol Biol. 12, 88 (1965). 32. Neidhardt, F. C , and Magasanik, B., Nature (London) 178, 801 (1956). 32a. Penninckx, M., Messenguy, F., and Wiame, J. M., Arch. Int. Physiol. Biochem 79, 204 (1971). 33. Ramos, F., Stalon, V., Piérard, A., and Wiame, J. M., Biochim. Biophys. Ada 139, 98 (1967). 34. Ramos, F., Thuriaux, P., Wiame, J. M., and Béchet, J., Eur. J. Biochem. 12, 40 (1970). 35. Salas, M. L., Salas, J., and Sols, A., Biochem. Biophys. Res. Commun. 31, 461 (1968). 36. Sheppard, D., and Englesberg, E., Cold Spring Harbor Symp. Quant. Biol. 31, 345 (1966). 37. Stachow, C. S., and Sanwal, B. D., Biochim. Biophys. Acta 139, 294 (1967). 38. Stalon, V., Ramos, F., Piérard, A., and Wiame, J. M., Biochim. Biophys. Ada 139, 91 (1967). 39. Thuriaux, P., Thesis, University of Brussels (1970). 40. Thuriaux, P., Ramos, F., Wiame, J. M., Grenson, M., and Bechet, J., Arch. Int. Physiol Biochem. 76, 955 (1968). 40a. Thuriaux, P., Ramos, F., Piérard, A., Grenson, M., and Wiame, J. M., in prepara­ tion. 41. Umbarger, H. E., Science 123, 848 (1956). 42. Umbarger, H. E., Annu. Rev. Biochem. 38, 323 (1969). 43. Umbarger, H. E., Curr. Top. Cell Regul 1, 57 (1969). 44- Wiame, J. M., in "Biochemical Evolution" (van Thoai and Roche, eds.), Gordon and Breach, New York, 1968. 45. Wiame, J. M., Rep. 10th Congr. MicrobioL, Mexico, 1970, in press. 46. Wood, W. A., Curr. Top. Cell. Regul. 1, 161 (1969). 47. Woolfolk, C. A., Shapiro, B., and Stadtman, E. R., Arch. Biochem. Biophys. 116, 177 (1966).

The Lac Repressor SUZANNE

I I |

BOURGEOIS

The Salk Institute for Biological Studies San Diego, California

I. Introduction 39 II. The Repressor Protein 42 A. Assays and Purification 42 B. Structure 44 C. General Properties 45 III. Inducer Binding 47 IV. Operator Binding 49 A. Equilibrium Studies 49 B. Kinetic Studies 54 C. The Structure of the Repressor-Operator Complex 57 V. The Mechanism of Induction 59 A. Interaction of Inducer and Anti-Inducer with the RepressorOperator Complex 59 B. Study of Repressore Altered by Mutations 61 VI. The Operator 63 A. Assay and Purification 63 B. Size and Properties 63 VII. Positive Control of the Lac Operon 65 VIII. The Mode of Action of the Lac Repressor 67 IX. Generalization 68 References 72

I. Introduction The lactose (lac) operon of Escherichia coli, its products and its regulation have been the subject of many reviews and of a recent comprehensive book (5). These have covered the extensive background of knowledge gathered over 25 years of in vivo studies mostly by Jacob and Monod and their co-workers and which led them to formulate in 1961 (36) their model of negative control of protein synthesis. This model has now been confirmed by in vitro experiments which permitted further analysis of the mechanism at the molecular level. The object of this review is to present and discuss the in vitro data which have recently become available on the molecular interactions which govern the expression of the lac operon. One should recall, however, the main features of the lac system and the conclusions regarding its regulatory switch, which were drawn from genetic and physiological studies and on which the in vitro analysis is based. 39

40

SUZANNE BOURGEOIS

PROMOTER-OPERATOR REGION REGULATORY GENE

STRUCTURAL GENES

P,0|

±mà i m-RNA 5' end POLYPEPTIDE CHAINS M . W . - 3 8 , 0 0 0

Lac mRNA 3'end -135,000

-30,000

NH2

-32,000 COOH

O

00

M TRANSPROTEIN ACETYLASE (SUBUNIT?) DIMER

REPRESS0R TETRAMER /3-GALACTOSIDASE TETRAMER

FIG. 1. Schematic representation of the lac region and its products. The small promoter-operator region (Section VI, B) is drawn out of scale. Arrows indicate the direction of transcription and of translation. For detailed information see reference (5).

The lac operon determines the synthesis of ß-galactosidase, the M protein which is an integral part of the galactoside permease, and the thiogalactoside transacetylase for which no physiological role is known. Regulation of the operon is revealed by an essentially coordinate synthesis of all three proteins, which is stimulated up to 1000-fold in the presence of inducing ß-galactosides. At the DNA level the lac operon consists of a cluster of the structural genes (z, y, and a), coding for those three proteins, adjacent to an operator (0) and a promoter (P), and transcribed into a single polygenic messenger RNA (Fig. 1). Transcription is very likely initiated at the promoter, although this point has not yet been clearly established in vitro. The distinction between the O and P regions and their size will be discussed in some detail later (Section VI, B). The regulatory gene, i, codes for the lac repressor (R) which, according to the Jacob-Monod model, blocks transcription of the operon by binding 0 while interaction of an inducer (I) with R prevents the R - 0 binding. The i gene is linked to the lac operon but is not part of it : it is transcribed separately but in the same direction as the operon (SI) and is under an independent control mechanism. The repressor is synthesized constitutively at the low level of

T H E LAC R E P R E S S O R

41

about 10 molecules per cell. Mutants (zq) have been obtained (59) which make more R than wild-type E. coli and indicate the existence of a promoter located at the operon-distal end of the i gene. The model was based on the isolation and characterization of a series of regulatory mutants, an approach which is now the classical way to unravel regulatory mechanisms of gene expression. The hydrolytic activity of one of the enzymes of the lac operon, the ß-galactosidase, was a fortunate feature which was exploited by the development of chromogenic substrates such as o-nitrophenyl-ß-galactoside to assay its activity or 5-bromo-3chloro-2-indolyl-ß-D-galactoside to detect constitutive strains, in addition to the usual indicator media for sugar utilization. More important, perhaps, was the introduction of nonmetabolized "gratuitous" inducers such as isopropyl-ß-D-galactoside ( I P T G ) . In parallel with the synthesis of ß-galactosides, a great virtuosity was developed in manipulating the lac region genetically. These genetic studies provide us today with the regula­ tory mutants needed for in vitro analysis as well as with transducing phages carrying the lac region (4) as a source of DNA about 70-fold enriched in operator (one operator per 30 X 106 daltons) as compared to the total E. coli chromosome (one operator per 2000 X 106 daltons). Constitutive mutants expressing a high level of operon activity in the absence of inducer defined the i gene and the O region. The recessive i~ mutations suggested the existence of R, the diffusible product of the trans-dominant i+ allele {65). The constitutive Oc mutations (87), cisdominant over the 0+ allele, pointed to O as the target of R action. How­ ever, the decisive criterion for negative control is the existence of iB (102) or superrepressed mutants with a lactose negative (lac~) phenotype dominant over the wild-type inducible one. This phenotype, which was correctly interpreted as the result of an impaired affinity of R for I, gave definitive proof for the direct role of R in repression by showing that R itself had a site for the binding of I as well as one for interacting with O. As for the nature of R, its interaction with O suggested that it might be RNA, while its specific interaction with I suggested that it was a protein. An argument in favor of the protein nature- of R was the finding of a thermosensitive repressor (86) stabilized by IPTG. On the other hand, some chloramphenicol inhibition experiments (66), since shown to have been misinterpreted (3, 34), argued against R being a protein. Solid evi­ dence that R was indeed a protein came from the suppression of i~ muta­ tions by well-characterized amber suppressors (9) whose mode of action at the level of translation had been precisely elucidated (16, 100). This was confirmed by a study (57) showing simultaneous reversion of i~ and non­ sense gal" cys~ mutations by unidentified suppressors. In contrast, none of the many constitutive O mutations tested were suppressible (9)—an argu-

42

SUZANNE BOURGEOIS

ment, negative as it is, against the translation of 0 . Further insight into the structure of R was gained by the finding of complementation between the products of some doubly mutated %H~ gene and an i+ allele (9). The very low frequency of this complementation phenomenon indicated that it was intracistronic resulting in a hybrid repressor made up of {iH~) {i+) subunit s conferring a superrepressed phenotype. This fact argued strongly that the i gene had only one cistron coding for a repressor made up of identical subunits. Similar complementing i~ mutants were later derived from i+, carefully mapped, and called i~d {i~~ dominant) {21). Altogether, on the basis of in vivo experiments, the main conclusions reached regarding the nature of the lac operon switch were that R is a polymeric protein capable of specific and mutually exclusive binding of O and I. Therefore the first in vitro data {29, 30) confirming that this was indeed the case brought no surprise, but great satisfaction. Surprises were, however, in store with regard to the kinetics of the R - 0 interaction {80) and the mechanism of induction {82), but those results had to await the development of a sensitive and convenient experimental methodology {8,78,81,84). II. The Repressor Protein A. Assays and Purification The first in vitro assay for R, developed by Gilbert and Müller-Hill {29) was based on the binding of the inducer I P T G measured by equilibrium dialysis. This assay allowed them to achieve a partial purification (to about 1% purity) using standard fractionation techniques, such as ammonium sulfate precipitation and DEAE chromatography, and to identify R as a protein of molecular weight about 150,000 to 200,000. Binding to the operator was detected by cosedimentation in a glycerol gradient of this repressor fraction, labeled in vivo with 35S, and of DNA from a lac trans­ ducing phage λφ80 dlac {30). Although these positive results were marginal, the controls fulfilled the expectations: binding was abolished by IPTG and weakened when the phage DNA carried a lac region altered by an Oc mutation. No binding was observed to the λφ80 helper phage DNA. The mere fact that binding could be detected at all under these conditions im­ plied that the binding had to be tight, with a binding constant on the order of ~10~ 12 M, in agreement with the value calculated by Koch {41) on the basis of in vivo experiments. Further characterization of R required its thorough purification as well as more convenient and sensitive binding assays. The major problem with the purification of R is its very low concentration in the cell, amounting to

THE LAC REPRESSOR

43

about 1/10,000 of the total proteins (see Section I ) . Two approaches have been utilized to get around this problem: mutants were selected that made more R (59) and purification procedures have been developed that take advantage of some rather unique properties of R (78). The combination of those two techniques makes the purification of R a fairly easy task today (58), although ending up with a fully active protein still poses some problems (see Section II, C). The rationale behind the purification procedure developed in our labora­ tory (78) is the fact that several enzymes interacting with DNA, such as DNA polymerase or exonuclease, were known to bind strongly to phosphocellulose, being eluted only at high salt concentration. Phosphocellulose chromatography turned out to be, indeed, the critical step in the purification, bringing about a 300-fold enrichment in R. This technique is now a classical step in the purification of a variety of DNA binding proteins such as RNA polymerase (14), where it allowed the detection of the sigma factor, the λ and 434 phages repressore (17) and the CAP (105) or CR (68) protein-mediating "catabolite repression" (see Sec­ tion VII). The new methodology developed in our laboratory to study the binding properties of R was based on the finding (78, 81) that R sticks to Millipore or membrane filters while retaining a bound ligand, either l4C-labeled I P T G or 32P-labeled λφ80 dlac DNA. Both the repressor-inducer complex (RI) and the repressor-operator complex (RO) trapped in the filter are stable enough to allow washings which essentially eliminate the background of free radioactive ligand. Since this filter technique appears to be a very general means for the study of ligand binding to proteins, it deserves some comments. Membrane filters had been used to retain nucleic acid-protein complexes such as ribosomes (63), and to study the binding of RNA polymerase to DNA (39) and of tRNA to amino acid synthetase (104) · They were originally thought to retain, specifically, nucleoprotein complexes under conditions where the protein by itself was not retained (39). How­ ever, Kuno and Kihara (44) have shown that it is a general property of proteins to be adsorbed on membrane filters. Hence the ligand need not be a nucleic acid. Retention is affected by ionic strength and magnesium is sometimes required. Different proteins not only require different ionic conditions for their retention to filters but some, such as L-lactic dehydrogenase, are inactive when bound to filters while others, such as polynucleotide phosphorylase (92) and RNA polymerase bound to DNA (2) retain their activity. Some alteration of proteins upon sticking to filters may stabilize the bound ligand against washing, and therefore need not be a drawback. This may be what happens in the case of the lac repressor

44

SUZANNE BOURGEOIS

and could explain why the RI complex is not readily dissociated upon washing of the filter in spite of the fact that binding of I P T G to R is not very tight with a dissociation constant Kd œ 10~6 M (see Section I I I ) . However, R is not inactivated upon its binding to filters in the form of RO complex, since the operator DNA (0) can be released by washing with IPTG, a property which was used to purify operator DNA {12) (see Section VI, A). Moreover, the filter does not seem to perturb the equilib­ rium of the R - 0 interaction {84) · The filter technique has since been used to study the binding of a variety of ligands to antibodies {20, 88) as well as the binding of lambdoid coliphages repressore {71,103) to their operators and of CAP to DNA (see Section VII). As applied to the lac repressor, it can detect as little as 10~16 mole of RO and its rapidity and convenience allowed the quantitative examination of the effect of a variety of experi­ mental conditions and ligands on the R - 0 interaction as well as the study of its kinetics (see Section IV). The detailed procedure and some useful modifications have been described {8). Other methods, more sensitive and convenient than equilibrium dialysis for measuring binding of I P T G to R, involve the precipitation of RI complex by a specific anti-R serum {78) or by ammonium sulfate {10). Some of the results obtained will be discussed in Section III. B. Structure Highly purified R sediments in a glycerol gradient as a 7 S component which cannot be distinguished from bovine 7-globulin by either sedimenta­ tion rate or chromatography on Sephadex G-200 {78), and which therefore must have a molecular weight close to 150,000. It can be dissociated by dodecyl sulfate {78) into ^ 3 S subunits of molecular weight ~40,000 indicating a tetrameric structure of R. More interesting is the fact that R dissociates spontaneously (i.e., for reasons still unclear) during purification {78) and upon storage {84) into 3 S subunits which have retained the capacity to bind I P T G but not O. This differential loss of activity will be discussed later (Section II, C). Amino acid analysis of the overproducing i q strain repressor {6) confirms that the subunits are identical, having a methionine amino terminus, a lysine carboxyl terminus, and a usual amino acid composition. It is an acidic protein with an isoelectric point of 5.6 which must, however, have a basic region binding to phosphocellulose at pH 7.0. No significant nucleic acid content could be detected by either ultraviolet absorption {78) or chemical analysis {58), but these results must be considered with caution since such analyses were performed on highly purified R preparations which had partially lost their O binding activity. The possibility of a cofactor involved in O binding will be dis­ cussed next (Section II, C).

45

THE LAC REPRESSOR

C. General Properties The lac repressor is involved in three specific binding interactions : R+4I^RI4

(1)

R + o ^± RO

(2)

RO + I — [IRO] ^± O + RI

(3)

Equations (1) and (2) describe the stoichiometry of the inducer and operator binding to R, the properties predicted by the Jacob-Monod model and discussed in detail in sections III and IV. Equation (3) represents the primary interaction responsible for the induction phenomenon as it was detected by kinetic studies (Section V, A) showing that I interacts directly with the RO complex, while binding of I to free R is only a secondary reaction. As mentioned before (Section II, B) the O binding activity of R decays preferentially to its I binding activity. This differential inactivation is accompanied by a dissociation of R into its I P T G binding subunits, although some inactivated tetramer is also seen (Fig. 2). This result indicates that the I P T G and 0 binding sites or R are distinct and suggests that the tetrameric form is necessary for tight O binding. It should be pointed out, however, that the failure to detect binding of O by the subunits may simply mean that the eventual binding was too weak to be detected under the conditions of our assay. Moreover these subunits have not so far been reassociated into tight 0 binding tetrameric R, but the fact that they can bind I P T G suggests that they may still be in a form close to their native state. On the other hand, the presence of differentially inactivated tetramer suggests that the tetrameric structure of R might not be sufficient for 0 binding and that a cofactor may be required. This cofactor, strongly bound to R, would be dissociated and lost during purification and storage. However, all attempts to detect such a cofactor have so far failed. The possibility, presented by Koch (42), that RNA might be associated with the R protein and mediate 0 binding seems unlikely because of the fact that O binding is insensitive to RNase (30). Pretreatment of R with high concentrations of EDTA does not affect its O binding activity, and Mg 2+ is the only element found so far to be necessary. The successful dissociation of the sigma factor from the core RNA polymerase on phosphocellulose (14) led us to try a similar experiment. However, when a purified R preparation of known O binding activity was submitted to phosphocellulose chromatography, its activity remained unchanged by the treatment (M. Clark, unpublished data). The genetic approach to the search for a repressor cofactor would be to select for constitutive mutations mapping outside of the lac region. Such mutants have never been observed, but, since the

46

SUZANNE BOURGEOIS

_

£ 80 3

-

1 ~\

(a)

7S

\

z s

•ΙΛ^ Z

5 60

O u

>— » >

-

ÎJ 40




o

Ö

>

Ö

o

Q

>

öd

o

Neurospora

Yeast

B. Fungi

Thiobacillus novellus

Yes

Yes

Yes

—

Yes

Relatively weak

None

None

AMP, ADP acti­ vates oxidative deamination and inhibits opposite direc­ tion. ATP antagonizes the AMP effect

None

Ammonia represses NAD enzyme and induces NADP enzyme. Glutamate has opposite effect

Ammonia represses NAD enzyme and induces NADP enzyme. Glutamate induces NAD enzyme

Glutamate induces NAD enzyme, re­ presses NADP enzyme. Arginine, asparate, and histidine repress NADP enzyme

Relative levels of two enzymes important in metabolic control

(cont'd.)

111-115

Relative levels of two 103-107 enzymes important in metabolic control

NAD enzyme involved 108-110 in a-KG synthesis. NADP enzyme in formation of glutamate

o

a

GO

o o

S3

»

Ü

> >

cl

O

Yes

Yes

Achyla Saprolegnia ► parasita Pythium debaryanum)

NADspecific

Blastocladiella emersomi

Organisms

No

No

NADPspecific

Presence of glutamate dehydrogenase

\\jominuea)

120,121

Regulation appears to be coupled with NADP specific isocitrate dehydro­ genase

Enzyme repressed by glucose and induced by glutamate

NADP, P-enolpyruvate, short-chain acyl-CoA deriva­ tives activate re­ ductive amination. Citrate and longchain acyl-CoA derivatives in­ hibitors

Reference

G T P and ATP activate re­ ductive amina­ tion. A M P inhibitor

Other comments

Regulatory patterns 116-119 appear to indicate direction enzyme operates sensitive to energy charge

Induction or repression

Ca 2+ and Mn 2+ acti­ vate reductive amination and in­ hibit the opposite direction. Isocitrate, fructose 1,6-diphosphate, fumarate inhibit oxidative deamination with no effect in opposite direction

Other effects

-ADIJIII 11

AMP and ADP activates. GDP, GTP, and ATP inhibit

Nucleotide effects

j

2

1—1

#

> to F

Ö Q

3

i>

3

t—1

o o

>

o

L-GLUTAMATE DEHYDROGENASES

107

for organisms which contain only a single glutamate dehydrogenase, it appears that in this organism the NAD enzyme serves primarily in a catabolic role (glutamate —> α-ketoglutarate) while the NADP enzyme is biosynthetic. Hollenberg et al. (107) have recently reported both glutamate dehydrogenases are located outside the mitochondria of yeast. In some organisms which have both an NAD- and NADP-specific enzyme, one of these may show allosteric properties and the function of the different enzymes in regulation may be more difficult to interpret. The chemoautroph Thiobacillus novellus is a specific example of this type of regulatory control. LéJohn and colleagues (108-110) have isolated the NAD- and NADP-specific glutamate dehydrogenases from this organism. They have observed that the latter is unaffected by purine nucleotides, but the former is specifically affected by AMP and, to a lesser extent, ADP. Although the effect of AMP is complex, initial velocity studies show that it activates α-ketoglutarate formation and inhibits glutamate formation. Thus, in the presence of AMP, the NAD-specific enzyme would utilize glutamate. However, ATP reverses the AMP effect and could therefore affect the direction of the reaction. Studies on the growth of the organism under different conditions (110) show that the NADP-specific enzyme is more prevalent when the organism is grown on thiosulfate, and less so when grown on glutamate. The NADP enzyme is also repressed by arginine, aspartate, and histidine. Fincham and Coddington (111, 112) isolated a number of mutant strains of Neurospora that require α-amino acids for normal growth. The mutants have been described as amination deficient due to their inability to incorporate inorganic nitrogen into amino acids. It was subsequently found that the mutants had a defective NADP-specific glutamate dehydro­ genase, although they did have a normal NAD-specific enzyme (113). Again, this implies that the NAD-specific enzyme functions in terms of glutamate formation. The reason these mutants grow poorly on inorganic nitrogen might arise from the fact that the conidia of wild-type organisms are deficient in the NAD enzyme (114) and this enzyme appears upon germination. Since the animation-deficient mutants have a defective NADP enzyme, there is no glutamate dehydrogenase in the conidia of these organisms. Sanwal and Latta (113) also suggested that if the two dehydrogenases are coupled, an effective transhydrogenase system exists. This can result in a regulation of the relative concentrations of pyridine nucleotide coenzymes. The levels of the two enzymes in Neurospora are regulated in a concerted way by the nature of the growth conditions. Thus, when Neurospora are grown on glutamate, the levels of the NAD enzyme are high and the NADPspecific glutamate dehydrogenase is repressed (115), but when ammonia is the nitrogen source, the levels of enzyme are reversed.

108

BARRY R. GOLDIN AND CARL FRIEDEN

The NADP-specific glutamate dehydrogenase from the various amination-deficient mutants of Neurospora form an interesting group of enzymes. Although these enzymes are inactive under conditions in which the wild-type glutamate dehydrogenase is active, some of the defective enzymes can be activated by brief heating or by preincubation with the same substances found to activate the wild-type enzyme. The activation effects are sigmoidal rather than hyperbolic and may reflect cooperative interactions. This obvious connection between the defective and wild-type enzyme has led to the suggestion that aberrant glutamate dehydrogenases have distorted allosteric interactions. Examples have been cited for organisms containing an NAD- or NADPspecific glutamate dehydrogenase, as well as those containing both enzymes. It has been pointed out that the NAD enzyme appears to act in a catabolic role while the NADP enzyme serves in an anabolic role. In organisms with two different enzymes, both an anabolic and catabolic role can be assigned to the glutamate dehydrogenase, and the relative rates of these two func­ tions are controlled by differential effects on the activity and synthesis of the two enzymes. Another type of situation can also prevail. In this case, both anabolic and catabolic processes are assigned to a single enzyme. While this requires a very complex glutamate dehydrogenase (like the animal enzymes), there is an example of a allosterically regulated NADspecific glutamate dehydrogenases found in the fungi of the Phycomycetes family in which there appears to be no NADP-specific enzyme. LéJohn and co-workers {116-121) have studied the NAD-specific glutamate dehydrogenase from a number of these organisms. There appear to be two classes of glutamate dehydrogenase which demonstrate different regulatory properties. Blastocladiella emersomi contains a gluta­ mate dehydrogenase whose activity is regulated by a number of substances in an interesting manner. Calcium and manganese activate the enzyme in the direction of the reductive deamination (117). This effect is most pro­ nounced at higher pH. In addition, isocitrate, citrate, fructose 1,6-diphosphate, and fumarate inhibit the oxidative deamination, but have no effect on glutamate formation (118). Blastocladiella also contains an NADPspecific isocitrate dehydrogenase which is activated by citrate. The com­ bination of complex regulatory effects has led to the proposal (118) that when the organism has an ample supply of energy (e.g., a high concentration of glucose), the mitochondria are hydrolyzing ATP with subsequent intake of Ca 2+ and release of H + and this forces glutamate formation. In addition, a buildup of compounds like citrate and FDP would activate isocitrate de­ hydrogenase thereby generating more a-ketoglutarate and also further forc­ ing the glutamate dehydrogenase in the direction of glutamate production by inhibiting the deamination reaction. This would create a glutamate sink.

L-GLUTAMATE DEHYDROGENASES

109

Since the isocitrate dehydrogenase reaction is NADP-specific and the gluta­ mate dehydrogenase reaction is NAD-specific an effective transhydrogenase system is also created which is of interest since this organism appears not to have a separate transhydrogenase (118). The Blastocladiella enzyme is also affected by a number of purine nucleotides. AMP and ADP activate the enzyme while GDP, GTP, and ATP inhibit (116). The activation by AMP arises from the rather complex kinetics demonstrated with respect to NAD. The substrate versus velocity curves have two plateaus and a mechanism for this behavior in terms of negative and positive homotropic interactions has been presented by Levitzki and Koshland (122). ATP is negative effector, thereby increasing the NAD concentration necessary for saturation while AMP is a positive effector giving rise to hyperbolic kinetics with respect to NAD. A number of other Phycomycetes have been studied by LéJohn and co-workers (120, 121). Included in this list are Achyla sp., Saprolegnia parasitica, and Pythium debaryanum. These organisms also have only the NAD-specific glutamate dehydrogenase, and this class of enzymes is affected by a number of compounds in a complicated manner. A total of five activators have been discovered. NADP, phospho-enolpyruvate, short-chain acyl-CoA derivatives, GTP and ATP act as unidirection activators of the biosynthetic reaction, while citrate, AMP, and long-chain acyl-CoA derivatives are inhibitors. To further complicate the situation, the activators antagonize the effects of the inhibitors. An explanation for this complex behavior has been proposed based on the coupling of the NADP-specific isocitrate and the NAD-glutamate dehydrogenase (121). Under energy-rich conditions when phosphoenolpyruvate, GTP, and ATP accumulate and activation of the biosynthetic reaction would result. The citric acid cycle would supply intermediates for amino acids and nucleotide synthesis. The coupling of isocitrate dehydro­ genase and glutamate dehydrogenase would result in a transhydrogenase in which NADPH would be synthesized at the expense of NADH when glutamate is being formed. This can be seen from the coupled reactions presented below: Isocitrate + NH 4 + + NADP+ + NADH — NAD+ + NADPH + glutamate + C0 2

The NADPH generated may be used in the synthesis of fatty acids. Activation by short-chain intermediates of fatty acid synthesis in a manner that favors NADH utilization fits this concept. Palmityl and oleyl-CoA which are end products of fatty acid synthesis, act as feedback inhibitors. Another interesting observation is that the distribution of the two complex NAD-specific glutamate dehydrogenase enzyme control systems among

110

BARRY R. GOLDIN AND CARL

FRIEDEN

the fungi is identical to the only two known pathways of lysine biosynthesis {121). The significance of this correlation is not known. The complex nature of the fungi enzymes, as pointed out previously, appears to be necessary to overcome the fact that these organisms only have the NAD-specific enzyme. In contrast to bacterial systems, which contain only one enzyme, the glutamate dehydrogenases in the fungi have to serve a more complex role. Thus, the added regulatory complexity. However, the fungi which have both glutamate dehydrogenases may also be complex. There have not been any detailed studies on the glutamate dehydrogenase in higher plants. Corn, pea, and oats {123,124) have been shown to contain an NAD-specific glutamate dehydrogenase associated with the mito­ chondria. In contrast, Vicia faba L. {125) can utilize both NAD and NADP. The NADP activity appears to be centered in the chloroplast, and the NAD activity in the mitochondria. Further work is required on purified glutamate dehydrogenases before any conclusions can be reached as to the predominant metabolic role this enzyme plays in higher plants. This survey of glutamate dehydrogenase from various sources indicates the diversity and complexity of this enzyme. It is clear, however, that the enzyme plays an important role in metabolic regulation. The type of enzyme present is controlled by the particular requirement of the organism with respect to growth conditions, life cycle, and energy requirements. Thus, the remarkable variation in the properties of the glutamate de­ hydrogenase are a reflection of the diversity between organisms with respect to the above-mentioned requirements and conditions. B. The Animal Mitochondria I Enzymes Previous sections have delineated the characteristics of glutamate dehydrogenase which are relevant to the control of enzymatic activity in animal tissues. These include, primarily, the fact that the enzyme can utilize both NAD(H) and NADP(H), that it is strongly affected by purine nucleotides, and that it undergoes a reversible polymerization reaction. There may be some question as to whether polymerization is an important control mechanism, but the fact that some kinetic characteristics do differ as a function of the extent of polymerization (see Section IV) and that the concentration of the enzyme in many tissues may be quite high ( > 2 mg/ml) certainly implies a physiological role for this process. It is of interest in this regard that different animal enzymes do differ in their ability to polymerize and that the different tissues of any given animal differ mark­ edly in the concentration of the enzyme in the mitochondria. For example, the concentration of the enzyme in skeletal muscle of various mammals is so low that probably no polymerization occurs even for an enzyme which

L-GLUTAMATE DEHYDROGENASES

111

would polymerize normally at high enzyme levels. On the other hand, the rat and dogfish liver enzymes do not appear to undergo any polymerization. In all cases, the enzyme does appear to be localized primarily in the mitochondria, despite some reports to the contrary. If one could extrapolate the data from nonanimal organisms to animal tissues, it might be possible to say that the dual coenzyme specificity of the animal glutamate dehydrogenases means that glutamate is formed using îsTADPH as the coenzyme and α-ketoglutarate is formed using NAD as coenzyme and that all the effects on activity, as discussed above, are designed to be able to control the utilization of one coenzyme relative to another as has been proposed (64-) · Although a large number of experiments, designed to determine the role of the enzyme have been performed, a unified picture for the metabolic role of glutamate dehydrogenase has not been achieved because of the complexity of the in vivo situation. Further­ more, a note of caution should be sounded regarding the conditions used in mitochondrial studies. Agents such as rotenone, dicumarol, dinitrophenol, inorganic phosphate, arsenite, ADP, and ATP are routinely added to mitochondrial preparations. Some of these compounds, e.g., rotenone, ADP, and ATP, have been shown to have direct effects on the activity of purified glutamate dehydrogenase. However, workers in this field have generally disregarded this problem. It should also be pointed out that uncoupling agents and respiratory chain inhibitors may cause large changes in the nature of the mitochondrial purine nucleotide pools, and in vitro studies indicate this type of alteration would also have a direct effect on glutamate dehydrogenase activity. These facts are generally ignored in the studies described below and the indirect effects of relating the ratio of purine nucleotides to pyridine nucleotides via energy linked transhydrogenation are emphasized. It is, however, instructive to cite briefly some of the conclusions drawn from studies with intact mitochondria. Borst (126) has presented data which indicate that the principal pathway for glutamate deamination in rat liver mitochondria is through the transamination reaction. Papa et al. (127) pointed out that both the transamination and deamination pathway are operative in mitochondria from liver and the relative importance of the two pathways depends on the state of the mitochondria. However, the transamination pathway appeared to be predominant despite the high levels of glutamate dehydrogenase present in liver mitochondria. This observation appears to demonstrate that the enzyme does not possess optimal activity in the mitochondria. In this regard, several workers (127-130) have observed that glutamate dehydro­ genase preferentially utilizes NADP(H) in mitochondria. This appears to be the case for the synthesis and breakdown of glutamate. This coenzyme selectivity has led to the suggestion (127, 128, 131) that NAD(H) is not

112

BARRY R. GOLDIN AND CARL FRIEDEN

available to glutamate dehydrogenase due to compartmentation. A further interesting rinding is that the deamination reaction is strongly inhibited when the NADPH:NADP ratio is high. This is true even though high concentrations of NAD are present in the mitochondria (132). Therefore the level of oxidation of NADP appears to be important in glutamate metabolism. In turn, this level of oxidation is dependent on the energy state of the mitochondria. Since these organelles possess an energy-de­ pendent transhydrogenase whose equilibrium is in the direction of NADPH production, the oxidation state of this coenzyme can be directly con­ trolled by energy levels in the mitochondria. Krebs and co-workers (188, 184), however, have arrived at a different conclusion regarding the coenzyme specificity of glutamate dehydrogenase. These investigators measured the concentrations of the oxidized and reduced substrates of glutamate dehydrogenase and ß-hydroxybutyrate dehydrogenase in rat livers frozen as soon as possible after death. The study was undertaken in an effort to determine the free NAD:NADH ratio in mitochondria, as calculated from the equilibrium constants of the respective dehydrogenase reactions and the experimentally determined substrate concentration in the cell. Results for the two dehydrogenases gave the same value for the NAD : NADH ratio implying that the reactions are in equilibrium with the same NAD-NADH pool. Further confirmation comes from the fact that the expression [NH3+][|S-hydroxybutyrate ][a;-ketoglutarate ] [acetoacet at e ] [glutamate ]

remains constant, as determined by measurement of substrate concen­ trations, in well fed, starved, and diabetic rats. If glutamate dehydrogenase were NADP specific via an energy-linked transhydrogenase system and in equilibrium with this couple, then the NAD:NADH ratio calculated from the glutamate dehydrogenase system should have been lower than the value calculated from the hydroxybutyrate dehydrogenase system (133). Furthermore, studies with rat livers deprived of blood for several minutes prior to freezing showed the same behavior as that described above (134)· Since the supply of higher-energy intermediates presumably ceases under these conditions, the mediation of an energy-linked transhydrogenase in the maintenance of the equilibrium between hydroxybutyrate dehydro­ genase and glutamate dehydrogenase is unlikely. Whether the glutamate dehydrogenase is active primarily with only one coenzyme, whether it is closely linked, via nucleotide effects, with the flux of substrates through the Krebs cycle, whether it serves to regulate the level of α-ketoglutarate, glutamate, ammonia, and/or coenzyme, or

L-GLUTAMATE DEHYDROGENASES

113

whether it performs several of these functions simultaneously are questions that have provoked considerable experimentation. In spite of in vivo studies and the wealth of information available from in vitro studies, it is still not possible to clearly define the metabolic role of this enzyme in animal tissue. There are several reasons for this dilemma. First, there are an almost bewildering number of compounds that affect activity in vitro, and effects depend upon which coenzyme is used and sometimes on the concentration of coenzyme. Second, the concentration of purine nucleotides, which are probably the most effective allosteric effectors, are not known within the mitochondria. Third, glutamate metabolic studies using liver slices, homogenates, or mitochondria are frequently complicated by the fact that purine nucleotide concentrations are not those which might exist in vivo. VIII. Conclusion An enormous amount of information is available for that class of enzyme known as glutamate dehydrogenase. As has been indicated throughout this chapter, the bovine liver has been most carefully characterized in terms of kinetic and molecular characteristics. The properties of this enzyme associated with the effects of purine nucleotides on activity and on the reversible association-dissociation reaction have served as models for the behavior of kinetically complex regulatory enzymes. The difficulties in evaluating the role of this enzyme in metabolic processes may arise from the fact that its role is markedly different in different tissues or under different circumstances within a given tissue. In order to indicate some generality to the enzyme, Frieden {64) has proposed that purine nucleotides may control the rate of utilization of one coenzyme relative to another. Within any given tissue, changes in purine nucleotide levels might then effect changes in coenzyme utilization. Even such a postulate, although relatively broad, may not be sufficient when comparing the metabolic function of different tissues. Certainly, one would not expect the enzyme to perform the same function in the liver, the muscle, and the brain. It is curious therefore that different tissues do not appear to possess different isozymes of the enzyme. This observation tempts us to speculate that the lack of isozymes and the complexity of the kinetic parameters are in fact related. The ability of the enzyme to utilize either NADH or NADPH, to catalyze a reaction with a number of different amino or keto acids, to be markedly affected by polymerization as well as by a variety of purine nucleotides as well as by some amino acids, phosphate, metal ions, and even (in high concentration) steroids or steroidlike substances, may reflect that, in fact, the enzyme may perform very different functions in different tissues without the availability of different isozymes.

114

BARRY R. GOLDIN AND CARL FRIEDEN REFERENCES

1. Frieden, C , in "The Enzymes" (P. D. Boyer, H. Lardy, and K. Myrbäck, eds.), Vol. 7, p. 3. Academic Press, New York, 1963. 2. Marier, E., and Tanford, C , J. Biol. Chem. 239, 4217 (1964). 8. Eisenberg, H., and Tomkins, G. M., / . Mol. Biol. 31, 37 (1968). 4. Appella, E., and Tomkins, G. M., J. Mol. Biol. 18, 77 (1966). 5. Smith, E. L., Landon, M., Piszkiewicz, D., Brattin, W. J., Langley, T. J., and Melamed, M. D., Proc. Nat. Acad. Sci. U. S. 67, 724 (1970). 6. Bitensky, M. W., Yielding, K. L., and Tomkins, G. M., / . Biol. Chem. 240, 1077 (1965). 7. Colman, R. F., and Frieden, C , J. Biol. Chem. 241, 3661 (1966). 8. Jornvall, H., Eur. J. Biochem. 16, 25 (1970). 9. Sund, H., and Burchard, W., Eur. J. Biochem. 6, 202 (1968). 10. Horne, R. W., and Greville, G. D., J. Mol. Biol. 6, 506 (1963). 11. Valentine, R. C , Abstr. 4th Eur. Regional Conf. Electron Microsc, Rome 2, 3 (1968). 12. Sund, H., Pilz, I., and Herbst, M., Eur. J. Biochem. 7, 517 (1969). 18. Eisenberg, H., in "Pyridine Dependent Nucleotide Dehydrogenases" (H. Sund, ed.), p. 293. Springer, Berlin, 1970. 14- Eisenberg, H., and Reisler, E., Biopolymers 9, 113 (1970). 15. Fisher, H. F., in " T h e Mechanism of Action of Dehydrogenases" (G. W. Schwert and A. D. Winer, eds.), p. 221. The Univ. Press, Kentucky, Lexington, 1969. 16. Reisler, E., Pouyet, J., and Eisenberg, H., Biochemistry 9, 3095 (1970). 17. Chun, P. W., Kim, S. J., Stanley, C. A., and Ackers, G. K., Biochemistry 8, 1625 (1969). 18. Dessen, P., and Pantaloni, D., Eur. J. Biochem. 8, 292 (1969). 18a. Cassman, M., and Schachman, H. K., Biochemistry 10, 1015 (1971). 19. Fisher, H. F., McGregor, L. L., and Power, U., Biochem. Biophys. Res. Commun. 8, 402 (1962). 20. Fisher, H. F., McGregor, L. L., and Cross, D. G., Biochim. Biophys. Acta 65, 175 (1962). 21. Cross, D. G., and Fisher, H. F., Arch. Biochem. Biophys. 110, 222 (1965). 22. Cross, D. G., and Fisher, H. F., Biochemistry 5, 880 (1966). 28. Fisher, H. F., and Bard, J. R., Biochem. Biophys. Res. Commun. 37, 581 (1969). 24. Fisher, H. F., and Bard, J. R., Biochim. Biophys. Acta 188, 168 (1969). 25. Huang, C , and Frieden, C , Proc. Nat. Acad. Sci. U. S. 64, 338 (1969). 26. Frieden, C , J. Biol. Chem. 238, 146 (1963). 27. Jirgensons, B., J. Amer. Chem. Soc. 83, 3161 (1961). 28. Bayley, P. M., and Radda, G. K , Biochem. J. 98, 105 (1966). 29. Sedgwick, K. A., and Frieden, C , Biochem. Biophys. Res. Commun. 32, 392 (1969). 30. King, K. S., and Frieden, C , J. Biol. Chem. 245, 4391 (1970). 31. Coffee, C. J., Bradshaw, R. A., and Frieden, C , unpublished results. 82. Frieden, C , Biochim. Biophys. Acta 62,421 (1962). 33. Anderson, P. J., and Johnson, P., Biochim. Biophys. Acta 181, 45 (1969). 84. Fahien, L. A., Wiggert, B. O., and Cohen, P. P., / . Biol. Chem. 240, 1083 (1965). 35. Corman, L., Prescott, L. M., and Kaplan, N . O., / . Biol. Chem. 242, 1383 (1967). 36. Barratt, R. W., and Strickland, W. N., Arch. Biochem. Biophys. 102, 66 (1963). 87. Barratt, R. W., Genetics 46, 849 (1961). 88. Winnocker, E. L., and Barker, H. A., Biochim. Biophys. Acta 212, 225 (1970). 39. Kaplan, N . O., Ciotti, M. M., and Stolzenbach, F . E., / . Biol. Chem. 221, 833 (1956).

L-GLTJTAMATE DEHYDROGENASES

115

40. Olson, J. A., and Anfinsen, C. B., / . Biol. Chem. 202, 841 (1953). 41. Fisher, H. F., and McGregor, L. L., Biochem. Biophys. Res. Commun. 34, 627 (1969). 42. Strecker, H. J., Arch. Biochem. Biophys. 46, 128 (1953). 43. Struck, J., and Sizer, I. W., Arch. Biochem. Biophys. 86, 260 (1960). 44· Caughey, W. S., Smiley, J. D., and Hellerman, L., / . Biol. Chem. 224, 591 (1957). 45. Bates, D . J., Goldin, B. R., and Frieden, C , Biochem. Biophys. Res. Commun. 39, 502 (1970). 46. Fisher, H. F., and McGregor, L. L., Biochem. Biophys. Res. Commun. 3, 629 (1960). 47. Frieden, C., J. Biol. Chem. 234, 809 (1959). 48. Wiggert, B. O., and Cohen, P. P., / . Biol. Chem. 241, 210 (1966). 49. Corman, L., and Kaplan, N . O., / . Biol. Chem. 242, 2840 (1967). 50. Engel, P . C , and Dalziel, K., Biochem. J. 115, 621 (1969). 51. Dalziel, K , and Engel, P. C., FEBS Letters 1, 349 (1968). 52. Cross, D . G., and Fisher, H. F., / . Biol. Chem. 245, 2612 (1970). 53. Pantaloni, D., and Dessen, P., Eur. J. Biochem. 11, 510 (1969). 54. Frieden, C , J. Biol. Chem. 234, 2891 (1959). 55. Fahien, L. A., and Stremcki, M., Arch. Biochem. Biophys. 130, 468 (1969). 56. Engel, P. C , and Dalziel, K., Biochem. J. 118, 409 (1970). 57. Hochreiter, M. C , and Schellenberg, K. A., / . Amer. Chem. Soc. 9, 6530 (1969). 58. Hochreiter, M. C , and Schellenberg, K. A., Fed. Froc. 29, 1215 (1970). 59. Fisher, H. F., Bard, J. R., and Prough, R. A., Biochem. Biophys. Res. Commun. 41, 601 (1970). 60. Bates, D. J., and Frieden, C , unpublished results. 61. Frieden, C , in "Role of Nucleo tides for the Function and Conformation of En­ zymes" (H. Kalckar, ed.), p. 194. Munksgaard, Copenhagen. 62. Iwatsubo, M., and Pantaloni, D., Bull. Soc. Chim. Biol. 49, 1563 (1967). 63. Huang, C , and Frieden, C , unpublished results. 64. Frieden, C , / . Biol. Chem. 238, 3286 (1963). 65. Colman, R. F., and Frieden, C , / . Biol, Chem. 241, 3652 (1966). 66. Frieden, C , and Colman, R. F., / . Biol. Chem. 242, 1705 (1967). 67. Monod, J., Wyman, J., and Changeux, J. P., / . Mol. Biol. 12, 88 (1965). 68. Jallon, J. M., diFranco, A., and Iwatsubo, M., Eur. J. Biochem. 13, 428 (1970). 69. Frieden, C , / . Biol. Chem. 245, 5788 (1970). 70. Frieden, C , / . Biol. Chem. 240, 2028 (1965). 71. Fahien, L. A., Wiggert, B. O., and Cohen, P . P., J. Biol. Chem. 240, 1091 (1965). 72. Yielding, K. L., Tomkins, G. M., Munday, J. S., and Curran, J., Biochem. Biophys. Res. Commun. 2, 303 (1960). 73. Yielding, K. L., and Tomkins, G. M., Proc. Nat. Acad. Sci. U. S. 46, 1483 (1960). 74. Tomkins, G. M., Yielding, K. L., and Curran, J., Proc. Nat. Acad. Sci. U. S. 47, 270 (1961). 75. Fisher, H. F., Cross, D. G., and McGregor, L. L., Nature (London) 196, 895 (1962). 76. Wolff, J., / . Biol. Chem. 237, 230 (1962). 77. Wolff, J., J. Biol. Chem. 237, 236 (1962). 78. Wolff, J., and Wolff, E. C , Biochim. Biophys. Ada 26, 387 (1957). 79. Yielding, K. L., and Tomkins, G. M., Proc. Nat. Acad. Sci. U. S. 47, 983 (1961). 80. Butow, R, A., Biochemistry 6, 1088 (1967). 81. diPrisco, G., Biochem. Biophys. Res. Commun. 26, 148 (1967). 82. Anderson, B. M., Anderson, C. D., and Churchich, J. E., Biochemistry 5, 2893 (1966).

116 83. 84. 85. 86.

BARRY R. GOLDIN AND CARL FRIEDEN

Piszikiewicz, D., Landon, M., and Smith, E. L., J. Biol. Chem. 245, 2622 (1970). Holbrook, J. J., and Jeckel, R., Biochem. J. I l l , 689 (1969). Goldin, B. R., and Frieden, C , Biochemistry. In press. Coffee, C. J., Bradshaw, R. A., Goldin, B. R., and Frieden, C , Biochemistry. In press. 87. Freedman, R. B., and Radda, G. K , Biochem. J. 114, 611 (1969). 88. Baker, J. P., and Rabin, B. R., Eur. J. Biochem. 11, 154 (1969). 89. Goldin, B. R., and Frieden, C , unpublished results. 90. Price, N. C , arid Radda, G. K , Biochem. J. 114, 419 (1969). 91. Hellerman, L., Schellenberg, K. A., and Reiss, O. K., J. Biol. Chem. 233, 1468 (1958). 92. Nishida, M., and Yielding, K. L., Fed. Proc. 29, 3792 (1970). 93. Bitensky, M. W., Yielding, K. L., and Tomkins, G. M., / . Biol. Chem. 240, 663 (1965). 94. Bitensky, M. W., Yielding, K. L., and Tomkins, G. M., / . Biol. Chem. 240, 668 (1965). 95. Nishida, M., and Yielding, K. L., Arch. Biochem. Biophys. 141, 409 (1971). 96. Malcolm, A. D. B., and Radda, G. K , Eur. J. Biochem. 15, 555 (1970). 97. Boezi, J. A., and DeMoss, R. D., Bacteriol. Proc. p. 124 (1959). 98. Goldfine, H., and Stadtman, E. R., / . Biol. Chem. 235, 2238 (1960). 99. Hardman, J. K , and Stadtman, T. C , / . Biol. Chem. 238, 2081 (1963). 100. Stern, J. R., and Bambers, G., Biochemistry, 5, 1113 (1966). 101. Halpern, Y. S., and Umbarger, H. E., J. Bacteriol. 80, 285 (1960). 102. Hooper, A. B., Hansen, J., and Bell, R., J. Biol. Chem. 242, 288 (1967). 103. Polakis, E. S., and Bartley, W., Biochem. J. 97, 284 (1965). 104. Bernhardt, W., Panten, K., and Holzer, H., Biochim. Biophys. Ada 99, 531 (1965). 105. Hölzer, H., Biochem. J. 98, 37P (1966). W6. DeCastro, I. N., Ugarte, M., Cano, A., and Mayor, F., Eur. J. Biochem. 16, 567 (1970). 107. Hollenberg, C. P., Riks, W. F., and Borst, P., Biochim. Biophys. Acta 201, 13 (1970). Î08. Lé John, H. B., Biochem. Biophys. Res. Commun. 28, 96 (1967). 109. LéJohn, H. B., Suzuki, I., and Wright, J. A., J. Biol. Chem. 243, 118 (1968). 110. LéJohn, H. B., and McCrea, B. E., / . Bacteriol. 95, 87 (1968). 111. Fincham, J. R. S., and Coddington, A., Cold Spring Harbor Symp. Quant. Biol. 28, 517 (1963). 112. Fincham, J. R. S., J. Biol. Chem. 182, 61 (1950). 113. Sanwal, B. D., and Lata, M., Can. J. Micro. 7, 319 (1961). 114- Sanwal, B. D., and Lata, M., Biochem. Biophys. Res. Commun. 6, 404 (1961). 115. Sanwal, B. D., and Lata, M., Arch. Biochem. Biophys. 97, 582 (1962). 116. LéJohn, H. B., and Jackson, S., J. Biol. Chem. 243, 3447 (1968). 117. LéJohn, H. B., Biochem. Biophys. Res. Commun. 32, 278 (1968). 118. LéJohn, H. B., / . Biol. Chem. 243, 5126 (1968). 119. LéJohn, H. B., Jackson, S., Klossen, G. R., and Sawula, R. V., J. Biol. Chem. 241, 5346 (1969). 120. LéJohn, H. B., and Stevenson, R., / . Biol Chem. 245, 3890 (1970). 121. LéJohn, H. B., Stevenson, R., and Meuser, R., / . Biol. Chem. 245, 5569 (1970). 122. Levitzki, A., and Koshland, D. E., Jr., Proc. Nat. Acad. Sci. U. S. 62, 1121 (1969). 123. Bone, D. H., Nature (London) 184, 990 (1959).

L-GLUTAMATE DEHYDROGENASES 124. 125. 126. 127.

117

Bulen, W. A., Arch. Biochem. Biophys. 62, 172 (1956). Leech, R. M., and Kirk, P. R., Biochem. Biophys. Res. Commun. 32, 685 (1968). Borst, P., Biochim. Biophys. Ada 57, 256 (1962). Papa, S., Palmieri, F., and Quagiariello, E., in "Regulation of Metabolic Process in Mitochondria" (J. M. Tager, S. Papa, E. Quagiariello, and E. C. Slater, eds.), p. 153. Elsevier, Amsterdam, 1966. 128. Papa, S., and Francavilla, A., in "Mitochondrial Structure and Compartmentation" (E. Quagiariello, S. Papa, E. C. Slater, and J. M. Tager, eds.), p. 363. Adriatica Editrice, Bari, 1969. 129. DeHaan, E. J., Tager, J. M., and Slater, E. C , Biochim. Biophys. Ada 131, 1 (1967). ISO. Papa, S., Tager, J. M., Francavilla, A., DeHaan, E. J., and Quagiariello, E., Biochim. Biophys. Ada 131, 14 (1967). 131. Papa, S., Tager, J. M., Francavilla, A., and Quagiariello, E., Biochim. Biophys. Ada 172, 20 (1969). 132. Tager, J. M., and Papa, S., Biochim. Biophys. Ada 99, 570 (1965). 133. Williamson, D. H., Lund, P., Krebs, H. A., Biochem. J. 103, 514 (1967). 134· Brosnan, J. T., Krebs, H. A., and Williamson, D. H., Biochem. J. 117, 91 (1970).

Regulation of Fatty Acid Biosynthesis* I

P. ROY VAGELOS

I I

Department of Biological Chemistry Washington University School of Medicine St. Louis, Missouri

|

I. Introduction II. Acetyl-CoA Carboxylase A. Structure-Function B. Control III. Fatty Acid Synthetase A. General Biosynthetic Scheme B. Acyl Carrier Protein C. Enzymes of Fatty Acid Synthetase of Escherichia coli D. Multienzyme Complexes E. Control References

119 120 123 133 141 141 143 146 150 159 162

I. Introduction The de novo synthesis of saturated fatty acids, studied in a variety of biological systems, is catalyzed by two enzyme systems, acetyl-CoA carboxylase and fatty acid synthetase. These enzyme complexes are found in the cell cytoplasm when cells are ruptured and the cellular components are fractionated by usual techniques. All the carbon atoms of the fatty acids are derived from acetyl-CoA, and Eq. (1) indicates the stoichiometry for the synthesis of palmitate from acetyl-CoA (85, 97). 8CH3CO—S—CoA + 7ATP + 14NADPH + 14H+ -> CH3(CH2)14COOH + 8C0A—SH + 7ADP + 7Pi + 14NADP+ + 6H 2 0

(1)

Although palmitate is the major fatty acid produced by most biosynthetic systems, the chain length of the fatty acid produced varies in different biological sources and under different experimental conditions. Examination of the substrates of fatty acid synthesis, acetyl-CoA, ATP, and NADPH, immediately suggests that this biosynthetic pathway can potentially be controlled by regulation of the availability of these substrates, and that is indeed the case. In mammals acetyl-CoA is produced within mitochondria, * The unpublished experimental work from this laboratory presented here and the preparation of this article have been assisted by grants from the National Institutes of Health (R01-HE-10406) and the National Science Foundation (GB-5142X). 119

120

P . ROY VAGELOS

and it apparently can reach the fatty acid biosynthetic enzymes in the cytoplasm only after conversion to citrate which is transported across the mitochondrial membranes to the cytoplasm (71, 144)· In the cytoplasm, citrate is cleaved by the citrate lyase to yield acetyl CoA. It has been shown that activity of the citrate lyase in liver and adipose tissue varies in parallel with fatty acid biosynthetic activity in a number of physiological condi­ tions, suggesting that this enzyme plays a role in regulating the supply of acetyl-CoA for fatty acid synthesis (78, 148, 158). The reductant for fatty acid synthesis is NADPH in most systems, and this is provided by both the pent ose phosphate pathway dehydrogenases and the "transhydrogenation cycle" enzymes in the cytoplasm. Glucose-6-P and 6-phosphogluconate dehydrogenases of the pent ose pathway produce NADPH which can be utilized in fatty acid synthesis directly (148, 150-153). In the "transhydrogenation cycle" oxaloacetate derived from citrate cleavage in the cytoplasm is reduced by malate dehydrogenase to malate which is con­ verted to pyruvate by malic enzyme. Pyruvate carboxylase, which is present in the cytoplasm of adipose tissue and liver, completes the cycle by catalyzing the carboxylation of pyruvate to form oxaloacetate (11, 37, 115, 129, 182, 186). The net result of this cycle is the conversion of NADH to NADPH at the expense of ATP. These reactions are demonstrated in Eqs. (2-5) : Oxaloacetate + NADH + H + - ^ malate + NAD+

(2)

Malate + NADP+ -> pyruvate + NADPH + H+ + C0 2

(3)

Pyruvate + ATP + C0 2 -► oxaloacetate + ADP + Pi

(4)

Sum: NADH + NADP+ + ATP -► NADPH + NAD + ADP + Pi

(5)

Activities of these enzymes have been shown to vary coordinately with acid biosynthesis in different physiological conditions, and they therefore been assigned regulatory roles and reviewed recently (148, In the present discussion only acetyl-CoA carboxylase and fatty synthetase will be considered in detail.

fatty have 153). acid

II. Acetyl-CoA Carboxylase Acetyl-CoA carboxylase catalyzes the first reaction in the synthesis of fatty acids which is unique to this biosynthetic pathway, and it is not surprising, therefore, that this enzyme is regulated. The discovery of the enzyme followed the observation by Klein (66) and by Gibson, Titchner, and Wakil (42) that bicarbonate is required in fatty acid synthesis. The report by Wakil, Titchner, and Gibson (174) that biotin is involved in this pathway was closely followed by the discovery of the enzyme by Wakil in 1958 (170).

R E G U L A T I O N O F FATTY ACID

121

BIOSYNTHESIS

Although acetyl-CoA carboxylase was the first biotin enzyme to be dis­ covered, the mechanism of action of biotin enzymes was elucidated by Lynen (85) and his co-workers in other biotin proteins. Studies of ß-methylcrotonyl-CoA carboxylase were the most productive to this end. Thus it was shown that this enzyme contains co valent ly bound biotin as a prosthetic group and that during the carboxylation a carboxybiotin enzyme inter­ mediate is formed (58, 69, 88). The reaction is demonstrated in Eqs. (6) and (7) : Mg2 +

ATP + HC03~ + biotin enzyme ;==± carboxybiotin enzyme + ADP + Pi

(6)

CH3

I

Carboxybiotin enzyme + CH3C=CHCO—S-^-CoA ;=± biotin enzyme CH2COO+ CH3C=CHCO—S—Co A

(7)

Avidin, a protein found in egg white, is a specific inhibitor of biotin, with which it forms a stoichiometric complex (34, 46, 177), and this inhibitor was used to illustrate the two partial reactions showrn in Eqs. (6) and (7). Lynen et al. (88) demonstrated an avidin-sensitive exchange reaction between ATP and either orthophosphate- 32 P or ADP-14C that required the presence of ATP, bicarbonate, ADP, Pi, Mg 2+ , and the active enzyme, thereby delineating the components of Eq. (6). The second partial reaction, shown in Eq. (7), wTas demonstrated as an avidin-sensitive exchange reac­ tion between 1,3,5-14C-labeled ß-methylglutaconyl-CoA and ß-methylcrotonyl-CoA. These exchange reactions led Lynen to postulate the exist­ ence of a carboxybiotin enzyme intermediate. The determination of the structure of the carboxybiotin enzyme inter­ mediate was facilitated by the discovery that ß-methylcrotonyl CoA carboxylase catalyzes the carboxylation of free ( + )-biotin: ATP + HCOr + (+)-biotin

► carboxybiotin + ADP + Pi l4

(8)

When this reaction was carried out with bicarbonate- C and high con­ centrations of ( + )-biotin, relatively large amounts of carboxybiotin-14C were produced. The labile carboxybiotin-14C was stabilized by methylation with diazomethane, and the methylated product was identified as V-Ncarboxymethylbiotin methyl ester by comparison with authentic com­ pounds (68, 88). The suspicion that the carboxylation of free ( + ) -biotin represented, as a model reaction, the carboxylation of the biotin prosthetic group of the enzyme was borne out by direct experimentation with the enzyme. Carboxybiotin enzyme, isolated from a reaction mixture containing

122

P. ROY VAGELOS

Lysine

FIG. 1. Chemical structure of carboxybiotin enzyme.

biotin enzyme, ATP, and bicarbonate-14C, was stabilized by esterification with diazomethane, and the methylated preparation was digested with pronase to yield l'-Af-carboxymethylbiocytin (69). Isolation of a biocytin derivative from the enzyme indicated that the biotin is bound to the enzyme protein through amide linkage at the e-amino group of a lysine residue. Figure 1 shows the chemical structure of carboxybiotin enzyme. Other biotin enzymes, which have also been shown to contain biocytin and to bind carbon dioxide in the same manner, include propionyl-CoA carboxylase (78), acetyl-CoA carboxylase (111), pyruvate carboxylase (136), and methylmalonyl CoA-oxaloacetate transcarboxylase (183). With each of the above enzymes it was possible to isolate the carboxybiotin enzyme in an active form and to demonstrate its ability to transfer the carboxyl group to an acceptor molecule as depicted in the two partial reactions of 0-methylcrotonyl-CoA carboxylase in Eqs. (6) and (7). The formation of carboxybiotin enzyme in the case of methylmalonyl-CoA-oxaloacetate transcarboxylase differs from the other biotin enzymes since methylmalonyl-CoA, rather than ATP and bicarbonate, donates the carboxyl group. Acetyl-CoA carboxylase, the enzyme involved in the synthesis of fatty acids, is a typical biotin enzyme. It catalyzes acetyl-CoA carboxylation according to the two partial reactions shown in Eq. 9 and 10 (103, 111, 169): Me*+

ATP + HC08~ + biotin enzyme ;=— carboxybiotin enzyme + ADP + Pi

(9)

Carboxybiotin enzyme + CH8CO—S—CoA ^ -O2CCH2CO—S—CoA + biotin enzyme

( 10)

Carboxybiotin enzyme has been isolated and characterized (111). Study of this enzyme in various biological systems has given new insight into the mechanism of action and regulation of biotin enzymes.

R E G U L A T I O N O F FATTY ACID

BIOSYNTHESIS

123

A. Structure-Function 1. MOLECULAR PROPERTIES OF ANIMAL ENZYME

In 1952 Brady and Gurin reported that long-chain fatty acid synthesis was greatly stimulated by citrate in a pigeon liver cell-free extract (18). Similar stimulation by certain tricarboxylic acid cycle intermediates, most notably citrate and isocitrate, was subsequently observed in experiments with homogenates of other animal tissues (1, 20, 75,121,123, HO). This has come to be known as the citrate effect even though isocitrate stimulates fatty acid synthesis as well as citrate in most systems (159). The stimula­ tory effect of citrate was localized to acetyl-CoA carboxylase simultaneously in three laboratories in 1962 (101, 102, 168). Studies of the citrate effect in rat adipose tissue acetyl-CoA carboxylase by Vagelos, Alberts, and Martin (160, 161) established that citrate is an allosteric activator of the enzyme, that activation of the enzyme by citrate is associated with the conversion of the inactive protomeric form to an active aggregated form, and that both the activation and the aggregation caused by citrate are freely reversible and parallel effects. These observations were later con­ firmed by studies of rat liver acetyl-CoA carboxylase by Matsuhashi, Matsuhashi, and Lynen (103) and Numa, Bortz, and Lynen (109), and by studies of avian liver enzyme in the laboratories of Lane (47, 50) and Numa (45, 113). Isolation of the avian liver (45, 47-50, 113), rat liver (106), and bovine adipose tissue enzymes (67) in homogeneous form has permitted their characterization, and the molecular properties are summarized in Table I. The avian liver enzyme has been studied most extensively in Lane's laboratory, where it has been shown to exist in two forms, an active poly­ meric form which has a molecular weight ranging from 4,000,000 to 8,000,000 and an inactive protomeric form with a molecular weight of 410,000 (47, 49, 50). These two forms have been investigated by electron microscopy, and Fig. 2 demonstrates the catalytically active polymeric form, which is seen in the characteristic filamentous form (aggregated form), and the catalytically inactive protomeric form (dissociated form). The filamentous polymeric form is 80-100 A in width by 0.2-0.5 μ in length, whereas the protomeric form appears particulate with dimensions of 80-140 Â. The polymeric forms are composed of 10-20 410,000 molecular weight protomers assembled in linear fashion, and they appear as filaments in the electron microscope. Factors that effect the reversible interconversion between the protomeric and polymeric forms have been investigated by Lane and his colleagues (47, 49, 50). Certain anions (citrate, isocitrate, malonate, tricarballylate, sulfate, and phosphate), acetyl-CoA, high protein concentration, and pH 6 to 7 promote aggregation of the protomer.

R a t liver

Citrate

Polymeric

Polymeric (filamentous) Protomeric

Citrate

Bovine adipose tissue

0 . 5 M NaCl, p H 8-9

Polymeric (filamentous) Protomeric Subunit

Form

Citrate, isocitrate or phosphate 0 . 5 M NaCl, p H 8-9 0 . 1 - 1 . 0 % SDS

Conditions

Avian liver

Source

TABLE I

51

13-15

106

— 57

— 550,000

14.4

67

Several million

— —

47, 49, 50

References

68

410,000 110,000

13.1 4.0

13-15 47-50

4-8 million

Molecular weight (sedimentation equilibrium)

55-59

Analytical ultra­ cent rifugati on (S)

47-50

Density gradient (S)

Sedimentation velocity

MOLECULAR P R O P E R T I E S O F A C E T Y L - C O A CARBOXYLASE

G F

< >

o

125

REGULATION OF FATTY ACID BIOSYNTHESIS

ÎÊÊÊM

m -*%i§^sft'?^irr

TilJNw

B

/

FIG. 2. Electron micrographs of avian liver acetyl-CoA carboxylase in (A) protomeric form and (B) polymeric form. Uranyl acetate stained X 120,000. From Gregolin et al. (47).

126

P. ROY VAGELOS

Dissociation of the polymer is caused by such factors as carboxylation of the enzyme to produce carboxybiotin enzyme, Cl~, and pH values greater than 7.5. Citrate and isocitrate, but not tricarballylate or phosphate, can promote the aggregation of the carboxybiotin enzyme to the polymeric state. In all reactions in which carboxybiotin enzyme is an intermediate citrate and isocitrate promote aggregation of the carboxylase to its poly­ meric form and cause a 15- to 16-fold activation of catalytic activity; neither phosphate nor tricarballylate has this effect. These studies have elegantly illustrated the striking correlation between state of activation and aggregation of acetyl-CoA carboxylase first demonstrated in the rat adipose tissue enzyme (160, 161). Homogeneous carboxylase has recently been isolated from bovine adipose tissue (67) and rat liver (106), and it can be seen in Table I that the molecular properties of these enzymes are very similar to those of the avian liver enzyme in that they exist in the active forms as large molecular weight aggregates. Binding studies, reported for the avian liver carboxylase (49), have revealed approximately one tight binding site for citrate (KD, 2 to 3 X 10~6 M) and one site for acetyl-CoA (KD, 4 X 10~6 M) per protomer (molecular weight 410,000). The biotin content, determined microbiologically with Lactobacillus arabinosus after acid hydrolysis of the protein, is 0.83 mole per protomer (50). The discovery of functional subunits in a bacterial acetyl-CoA car­ boxylase stimulated an examination of the subunit structure of the avian liver enzyme (49). Dissociation was achieved with 1% sodium dodecyl sulfate which gives rise to subunits with a sedimentation coefficient (s2o,w) of 4.3 S. Subunit molecular weight determined by sedimentation equilib­ rium, corrected for detergent binding, is 114,000 and by electrophoresis in SDS-polyacrylamide gels, is 110,000. These subunits are nonidentical, as indicated by the fact that there is a single biotin prosthetic group per 410,000 molecular weight protomer. However, it has not yet been possible to determine the functions of the individual subunits since they are de­ natured by the SDS required for dissociation. 2. SUBUNIT FUNCTIONS

The demonstration that the animal acetyl-CoA carboxylase contains protomeric units of 410,000 molecular weight with a single biotin prosthetic group suggested that this enzyme might be analogous to the animal fatty acid synthetase. The latter enzyme system is a very tightly associated multienzyme complex with a molecular weight of approximately 450,000 daltons, and it contains 1 mole of prosthetic group per mole of enzyme complex. In bacteria and plants, enzymes which comprise the fatty acid synthetase are found completely dissociated when cells are ruptured, and

127

REGULATION OF FATTY ACID BIOSYNTHESIS

therefore it has been possible to identify and study all the component proteins of the synthetases in these dissociated systems. For this reason extracts of Escherichia coli were examined for acetyl-CoA carboxylase with the hope that active subunits could be isolated (9). Purification of the carboxylase by ammonium sulfate fractionation followed by adsorption and elution from alumina C7 led to a complete loss of activity which, however, could be reconstituted by mixing two protein fractions obtained from the alumina C7 step. The requirement for these two fractions, E a and Eb, for acetyl-CoA carboxylase activity is demonstrated in Table II, which also shows the requirements for Mn2+, ATP, and acetyl-CoA. Using the conversion of bicarbonate-14C to malonyl-14C CoA as an assay, E a and E b were each highly purified so that they could be examined for biotin and tested in the partial reactions shown in Eqs. (9) and (10). As noted in Table III, acid hydrolysis of the proteins followed by microbiological assay indicated that E a contained 2.3 nmoles of biotin per milligram of protein, whereas E b contained no biotin. Each protein was then incubated with ATP, bi­ carbonate- 1 ^ and MnCl2, and the reaction mixtures were filtered through Sephadex G-50 to separate the proteins from the low molecular weight radioactive compounds. As noted in Table III, 2.4 nmoles of 14C-carboxy-Ea was formed under these conditions, an amount equivalent to the amount of biotin in E a . On the other hand, Eb, which contained no biotin, contained no radioactivity. TABLE II REQUIREMENTS FOR MALONYL-COA FORMATION

Components Complete system0 -Ea -Eb -MnCl 2 -ATP -Acetyl-CoA

Malonyl-CoA (nmoles) 3.7 0 0 0 0.28 0

° The complete reaction mixtures contained 55 mili imidazole-HCl buffer, pH 6.7; 0.44 mikf MnCl2; 0.44 milf ATP; 0.3 m l acetylCoA; 14 m l KH 14 C0 3 (0.2 MCi/Mmole) ; 0.005 unit each of E a and E b in a total volume of 0.09 ml.

128

P. ROY VAGELOS TABLE I I I B I O T I N CONTENT AND FORMATION OF E - 1 4 C 0 2

Enzyme Ea Eb

Biotin content (nmoles/mg protein)

E- 1 4 C0 2 " formed (nmoles/mg protein)

2.32 0

2.42 0

° Biotin content was determined microbiologically. E-14CC>2~ formation was determined in the following system: imidazole-HCl, p H 7.5; LO mg E a or E b ; 0.5 m l MnCl 2 ; 0.5 m l A T P ; and 1.0 m l NaH 1 4 C03 (20 mCi/inmole). After 2 minutes incubation at 25, mixtures were filtered through Sephadex G-50.

Further studies indicated that the formation of E a - 14 C0 2 ~ was inde­ pendent of E b ; addition of E b to the carboxylation mixtures neither in­ creased nor decreased the amount of carboxybiotin-E a . Thus it was con­ cluded that E a functioned in the reaction of Eq. (9), where carboxy­ biotin-Ea is the product of the reaction. With isolated 14 C-carboxybiotin-E a available, the function of Eb was investigated in the second partial reaction, indicated in Eq. (10). Table IV demonstrates that E a - l4 C0 2 ~ is efficiently converted to malonyl-l4C-CoA and that this conversion requires acetyl-CoA and the second protein fraction, E b . These experiments indicated that E. coli acetyl-CoA carboxylase is easily dissociated into two components: E a which contains biotin and which functions in the first partial reaction in which carboxybiotin-E a is formed, Eq. (9) ; and E b , which is required T A B L E IV REQUIREMENTS FOR CARBOXYL T R A N S F E R FROM E a - 1 4 C 0 2 _ TO A C E T Y L - C O A 14 14

System 0

Ea- COr (nmole)

Complete Complete Complete -Eb — Acetyl-CoA

0.0055 0.013 0.037 0.037 0.037

C-Malonyl-CoA formed (nmole) 0.0052 0.011 0.021 0 0

Yield

(%) 94.7 84.5 56.6 0 0

° The complete system contained 55 m l imidazole-HCl, p H 7.5; 0.15 m M acetyl-CoA; 0.003 unit E b ; and E a - 1 4 C0 2 ~ in the amounts indicated in a volume of 0.09 ml. Reactions were incubated for 1 minute, and the malonyl-CoA was determined as acid stable radioactivity.

129

REGULATION OF FATTY ACID BIOSYNTHESIS

for carboxyl transfer from carboxybiotin-E a to acetyl-CoA to form malonylCoA, Eq. (10). Further studies of E a were greatly hampered by the extreme lability of this fraction. This problem was solved by the discovery that E a is quite stable when maintained in 20% glycerol (8). To facilitate further in­ vestigations, three different assays were utilized to measure E a . In the first of these, acetyl-CoA carboxylation was measured in the presence of excess Et>; this represented the overall reaction. Second, it was discovered that E a catalyzes the carboxylation of free ( + )-biotin, the reaction indi­ cated in Eq. 8. Catalysis of free ( + )-biotin carboxylation by the ß-methylcrotonyl-CoA carboxylase (68, 88) and by avian liver acetyl-CoA carboxylase (146) had been reported earlier. The third assay utilized a biotin auxotroph of E. coli which was grown in the presence of biotin-14C so that E a , containing covalently bound biotin-14C, could be followed even under conditions where catalytic activity was lost. Figure 3 shows the results of a sucrose density gradient experiment in which biotin- 14 C-E a was centrifuged

pH 78, 15 Hours

Biotin

Carboxylase

1

100>^>

Acetyl-CoA Carboxylase 50

0

10 FRACTION

20 NUMBER

30

FIG. 3. Sucrose density gradient of 14C-biotin-Ea at pH 7.8. 14C-biotin-Ea (0.1 mg, 1000 cpm, 0.08 unit acetyl-CoA carboxylase, 0.15 unit biotin carboxylase) in 100 μΐ 0.02 M potassium phosphate buffer and 20% glycerol was layered over a 5-20% sucrose gradient containing 0.02 M potassium phosphate buffer pH 7.8, and 20% glycerol and centrifuged for 15 hours at 5°. Aliquots were counted (O—O), assayed for acetyl-CoA carboxylase in the presence of E b (A—A), and biotin carboxylase ( · — # ) .

130

P. ROY VAGELOS

pH 90 21 Hours

800

C cpm

\ \ Biotin

Carboxylase

6.9S

11.2 S

1

i

H600

X

400 I

200

10 20 FRACTION NUMBER

FIG. 4. Sucrose density gradient centrifugation of biotin-14C-Ea at pH 9.O. Procedure was the same as in experiment of Fig. 3 except that buffer was 0.02 M Tris-HCl, pH 9.O.

for 15 hours at pH 7.8. It is noted that E a activity, as measured in the acetyl-CoA carboxylase assay with excess Eb, was approximately in the middle of the gradient and that free ( + )-biotin carboxylase activity sedimented with a peak in the same region. In addition there was a radio­ active peak, as expected, in the same location indicating the presence of protein-bound biotin-14C in E a . Completely unexpected was the presence of a second, much more slowly sedimenting peak of biotin-14C. The finding of this slower-sedimenting biotin-14C peak suggested that E a partially dissociated under those experimental conditions to give rise to a low molecular weight subunit which contained the covalently bound biotin. Various conditions were tested in attempts to completely dissociate E a into its components, and the experiment of Fig. 4 shows that centrifugation of biotin- 14 C-E a at pH 9.0 allowed total dissociation of E a into two com­ ponents: one of these contained essentially all the covalently bound biotin-14C, had a sedimentation coefficient of 1.3 S, and had no catalytic activity in the assays tested. The second component catalyzed free ( + )-biotin carboxylation and had a sedimentation coefficient of 5.4 S, but it contained very little biotin-X4C. These experiments indicated that E a dissociates into two subunits, the catalytic unit, called biotin carboxylase

R E G U L A T I O N O F FATTY ACID

BIOSYNTHESIS

131

(BC), and the biotin-containing unit, called biotin carboxyl carrier protein (BCCP). The functions of the three subunits, BC, BCCP, and E b , in the acetyl-CoA carboxylase reaction are summarized in the reactions of Eqs. (11) and (12). BC, Mn2 +

ATP + HCO3- + BCCP

- CO2-—BCCP + ADP + Pi

(11)

Eb

CO2-—BCCP + CH3CO—S—CoA ^± -02CCH2CO—S—CoA + BCCP

(12)

The biotin protein, BCCP, has been obtained pure in at least three different homogeneous forms from extracts of E. coli (107). The largest of these has a molecular weight of approximately 40,000, and, like the smaller forms, is free of BC and Eb activities. Only small quantities of this form have thus far been isolated. However, preliminary experiments indi­ cate that it is significantly more active than the smaller forms of BCCP, exhibiting a higher Vm&* and lower Km in the acetyl-CoA carboxylase reaction. This larger form of BCCP spontaneously dissociates to lower molecular weight fragments. A second form of BCCP, obtained when the larger form dissociates, has a molecular weight of 10,267 and corresponds to the 1.3 S sedimenting protein demonstrated in Fig. 4. The amino acid composition of this protein, which contains one biotin per mole of protein, is indicated in Table V, which also lists the composition of BCCP obtained from a different biotin enzyme, methylmalonyl-CoA-oxaloacetate transcarboxylase, of Propionibacterium shermanii. A third form of BCCP has been isolated and crystallized. This form has a molecular weight of 9100, and amino acid analyses reveal that it contains 14 amino acids less than the 10,267 molecular weight species; therefore it is assumed that the 9100 molecular weight form is derived from the larger peptide as the result of proteolytic cleavage. Enzyme kinetic measurements indicate that the Km (2.5 X IO -5 M) and F m a x (0.2 Mmole/min/mg protein) are exactly the same for the two smaller forms of BCCP in both the biotin carboxylase reaction (Eq. 11) and the overall acetyl-CoA carboxylase reaction. The precise relationship between the 40,000 molecular weight BCCP and the small forms is not understood. It is possible that the large form is composed of aggregates of the smaller BCCP or a single BCCP plus a nonbiotin peptide. The other two subunits of acetyl-CoA carboxylase, BC and E b , have been studied to a lesser extent. BC has recently been obtained in homo­ geneous form and crystallized (32). BC has a molecular weight of ap­ proximately 100,000 and is composed of two 50,000 dalton subunits. Of particular interest are the relative affinities of this subunit for free ( + )biotin and for the biotin protein, BCCP. The small forms of BCCP have an apparent Km of 2.5 X 10~5 M in the reaction of Eq. (11), whereas free

132

P.

ROY

VAGELOS

TABLE V AMINO ACID COMPOSITION OF B C C P

Amino acid Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half-cystine Valine Methionine Isoleu cine Leu cine Tyrosine Phenylalanine Total M.W. a

Escherichia coli: Acetyl-CoA carboxylase (residues/mole)

Propionibacterium shermanii: methylmalonyl-CoA oxaloacetate transcarboxylase" (residues/mole)

5 1 2 6 5 6 13 8 6 12 1 12 5 6 3 1 3

8 2 2 8 5 4 8 12 11 9 1 9 2 5 6 2 3

95 10,267

97 11,000

From Gerwin et al. (39).

( + )-biotin has an apparent Km of 8.4 X IO -2 M, an impressive demon­ stration that the peptide structure of BCCP is very useful in biotin carboxylation (107). The third protein of acetyl-CoA carboxylase, E b , is specifically involved in the transfer of the carboxyl group of C0 2 ~-BCCP to the acceptor, acetyl-CoA. Preliminary experiments indicate that E b contains the binding site for acetyl-CoA. Recent reports of Wood and his co-workers (3, 39, 64) have indicated that methylmalonyl-CoA-oxaloacetate transcarboxylase, another biotin enzyme, has also been dissociated into nonidentical subunits. One of the subunit s has a sedimentation coefficient of 1.3 S and contains the amino acids listed on Table V and approximately 1 mole of biotin. Although the roles of the other subunits of this enzyme have not been defined, it is ap­ parent that the biotin containing subunit is comparable to BCCP of

R E G U L A T I O N O F FATTY ACID

BIOSYNTHESIS

133

acetyl-CoA carboxylase. In light of these findings it is tempting to postulate that all biotin enzymes might be composed of at least three different subunits; a biotin protein such as BCCP, a biotin carboxylase, and a subunit specifying the acceptor molecule. I t is also possible that various biotin enzymes in the same organism will contain similar BCCP and biotin carboxylase subunits and will differ from each other only in containing different E b subunits which confer acceptor specificity on the enzyme complexes. The biotin protein, BCCP, in many features resembles the acyl carrier protein, ACP, which is a component of the fatty acid synthetase. Both have a molecular weight of approximately 10,000 daltons. Both are com­ ponents of complex enzyme systems which are readily dissociable in E. coli but firmly associated in other organisms. Both have covalently bound prosthetic groups, biotin in BCCP and 4'-phosphopantetheine in ACP. The prosthetic groups of both proteins function in their respective reactions as covalent substrate binding sites. Because of these analogies, the biotin protein is considered to be a carboxyl carrier protein. It is obvious that major differences exist between the bacterial biotin enzymes, which are easily dissociated into active subunits, and the animal acetyl-CoA carboxylase, which can be dissociated only under denaturing conditions. An intermediate form of acetyl-CoA carboxylase has been studied in wheat germ by Heinstein and Stumpf (55). This enzyme system has been dissociated into two active protein components: one of these has a sedimentation coefficient of 7.3 S and corresponds to E b , and the other has a sedimentation coefficient of 9.4 S and corresponds to E a of the E. coli enzyme. Further resolution to separate the BCCP and biotin car­ boxylase from the E a fraction has not yet been possible. B. Control 1. ALLOSTERIC REGULATION

a. Citrate Effect. Citrate (isocitrate) activation of acetyl-CoA carboxylase has been studied in detail in several laboratories (47, 101, 105, 109, 159, 161). As mentioned above, activation is associated with polymerization of inactive protomers to form active polymeric filaments. Kinetic analyses of the enzyme of rat liver (112), avian liver (48, 131), and bovine adipose tissue (74) have established that the major effect of tricarboxylic acid activators is on the maximal velocity of the enzyme rather than on the Km s for substrates. Thus experiments have been aimed at discovering how conformational changes at the active site, induced by the binding of citrate at the allosteric site, result in the increased efficiency of the enzyme. In 1964, Matsuhashi, Matsuhashi, and Lynen (103) attempted to

134

P. ROY VAGELOS

localize the site of citrate activation by studying the two partial reactions in the rat liver enzyme. Using the exchange reactions between ATP and either orthophosphate-32P or ADP-14C, which represent the first partial reaction (Eq. 9), and the exchange reaction between malonyl CoA and acetyl-14C-CoA, which represents the second partial reaction (Eq. 10), it was found that citrate activated both partial reactions. Similar results with exchange reactions were obtained with the avian liver enzyme {48, 50,131). Further studies of the partial reactions in Lane's laboratory have been facilitated by the use of model compounds. Careful kinetic analyses of the partial reactions is impossible due to the extremely rapid rates observed when substrate amounts of enzyme are utilized. As mentioned earlier, acetyl-CoA carboxylase catalyzes the carboxylation of free ( + )-biotin. Since this carboxylation is a model for the first half reaction, and ( + ) -biotin carboxylation proceeds much more slowly than the carboxylation of the biotin enzyme, the effect of citrate could be tested directly {145). As anticipated, citrate greatly stimulated the carboxylation of ( + )-biotin; however, it also caused an increase in the Km for ( + ) -biotin. This has been interpreted as suggesting that citrate causes tighter binding of the biotin prosthetic group, thereby making it more difficult for the ( + )-biotin to enter the carboxylation site. Kinetic analysis of the second partial reaction, that between carboxybiotin enzyme and acetyl-CoA, is also impossible due to extremely rapid rates, and this has been solved by using acetyl pantetheine as the carboxyl acceptor. Acetyl pantetheine is a model for acetyl-CoA, and the carboxylation of this model compound by carboxy­ biotin enzyme is slow enough so that kinetic studies of the effect of citrate have been possible. As anticipated from the exchange reaction studies, citrate (isocitrate) greatly stimulates the carboxylation of acetyl pante­ theine by carboxybiotin enzyme. Thus all the above studies indicate that both partial reactions are stimulated by citrate. A component of the enzyme involved in both half reactions is the biotin prosthetic group, and therefore Lane has attempted to obtain evidence that citrate causes conformational changes in the vicinity of the protein-bound biotin which are associated with enhanced reactivity of the carboxylated biotin prosthetic group. Such evidence has been obtained from two types of experiments. In the first of these {181) avidin, the specific biotin inhibitor, was incubated with the enzyme under conditions in which acetyl-CoA carboxylase was completely inactivated in less than 1 minute. The presence of citrate during the incubation of enzyme with avidin completely pre­ vented the inactivation of the enzyme by the avidin. Thus citrate induces conformational changes in the vicinity of the biotin prosthetic group which prevent the access of avidin to the prosthetic group.

REGULATION OF FATTY ACID BIOSYNTHESIS

135

A second group of experiments have attempted to demonstrate the effect of citrate on the reactivity of the carboxylated biotin prosthetic group (131). In this instance, the effect of citrate (isocitrate) on the rate of decarboxylation of malonyl-1,3- l4 C-CoA, catalyzed by the enzyme, and on the rate of decarboxylation of carboxybiotin enzyme were examined. It was found that malonyl-CoA decarboxylation was greatly stimulated by citrate when acetyl-CoA was present, indicating that both citrate and acetyl-CoA effect the reactivity of carboxybiotin enzyme, which is an intermediate in malonyl-CoA decarboxylation. Carboxybiotin enzyme was also tested directly, and it was found that citrate greatly stimulates decarboxylation of the enzyme, especially when acetyl-CoA is present. These experiments suggest that citrate in the presence of acetyl-CoA increases the susceptibility of carboxybiotin enzyme to decarboxylation, presumably due to conformational changes in the vicinity of the biotin prosthetic group. Animal acetyl-CoA carboxylase can exist as a catalytically inactive protomeric or a catalytically active polymeric form, and the activity of the enzyme is determined by the position of the protomer-polymer equilibrium. Citrate, presumably by binding preferentially to the polymeric form, can shift the equilibrium toward the active polymeric form. Citrate activation is associated with conformational changes at the active site in the vicinity of the biotin prosthetic group, as indicated by the decreased reactivity of the prosthetic group with avidin and by the increased reac­ tivity of carboxybiotin enzyme brought about by citrate (isocitrate). Such allosteric effects of tricarboxylic acids have not been noted in the bacterial (8, 9) or plant (55) acetyl-CoA carboxylases studied thus far. ò. Feedback Inhibition. Regulation of saturated fatty acid biosynthesis was reviewed recently, and the possibility of feedback inhibition of acetylCoA carboxylase was discussed (97). The observation that fatty acid synthesis is decreased in starvation and diabetes and stimulated by refeeding and insulin treatment could be explained by connecting two im­ portant observations. The first was that acetyl-CoA carboxylase is inhibited by palmityl-CoA and other long-chain acyl-CoA's (16, 109). The inhibition is competitive with regard to citrate, an allosteric activator of the enzyme, but noncompetitive with regard to substrates, acetyl-CoA, bicarbonate, or ATP. The Ki for palmityl-CoA is approximately 0.8 μΜ. Centrifugation studies showed that palmityl-CoA, which inhibits the enzyme competi­ tively with citrate, caused the enzyme to dissociate into its protomeric state. The second observation was that long-chain acyl-CoA derivatives were found in increased concentrations in livers of starved and fat-fed rats (17, 156). Since increase in liver fatty acyl-CoA's was presumed to be

136

P. ROY VAGELOS

secondary to adipose tissue lipolysis, it was reasonable to suspect that various physiological conditions associated with increased lipolysis and decreased fatty acid synthesis might be explained by inhibition of acetylCoA carboxylase by long-chain acyl-CoA derivatives in the liver. This proposal has been questioned because long-chain acyl-CoA's are potent detergents which inhibit many enzymes nonspecifically. Taketa and Pogell (149) demonstrated inhibition of a variety of enzymes, including glutamate, malate, glycerol-3-P, isocitrate, 6-P-gluconate, and glucose-6-P dehydrogenases, as well as fumarate, by palmityl-CoA. In view of the widespread inhibition by palmityl-CoA and other thioesters, the physiological role of these esters as regulators of certain metabolic pathways was seriously questioned. An important fact, in considering a physiological role for thioesters of Co A in inhibiting acetyl-CoA carboxylase and other enzymes, is that serum albumin prevents the inhibitory action of palmityl-CoA on many enzymes, probably because serum albumin binds the thioesters very strongly (109, 149). Since palmityl-CoA binds to so many proteins, how likely is it that there would be a high enough concentration of free thioesters in the liver to inhibit acetyl-CoA carboxylase? Inhibition by palmityl-CoA of citrate synthetase {143, 155, 178) and fatty acid synthetase (33, 79) have also been observed, with similar argu­ ments as above suggesting that these inhibitions have no physiological regulatory significance (33, 143). One approach that would support a physiological role for long-chain thioesters in regulation of fatty acid synthesis might be the demonstration of inactive (inhibited) acetyl-CoA carboxylase in liver extracts of starved or diabetic animals. Reactivation of the enzyme by removal of inhibitor (Sephadex filtration or dialysis) would certainly support the proposed role of acyl-CoA thioesters if the inhibitor were a thioester. However, attempts to identify thioester-inhibited carboxylase have been negative. Izui and his co-workers (61) have recently attempted to assign a physio­ logical regulatory role for free fatty acids or their CoA derivatives in E. coli metabolism since they have shown that these compounds greatly stimulate the activity of phosphoenolpyruvate carboxylase in vitro. Although these authors demonstrated that the activation by these compounds is allosteric in nature, and that the fatty acid or fatty acyl-CoA activator site is differ­ ent from the acetyl-CoA allosteric site of this enzyme, the argument for a physiological role of the fatty acyl-CoA as an activator of this enzyme is weakened by the fact that the enzyme is also equally stimulated by longchain alcohols, such as ?i-heptyl alcohol, and by dioxane. A mutant which had lost the sensitivity to fatty acids and long-chain thioesters also lost the sensitivity to organic solvents, suggesting that enzyme stimulation by hydrophobic compounds may be nonspecific. Thus a definite physiological

REGULATION OF FATTY ACID BIOSYNTHESIS

137

role for long-chain acyl-CoA's has not been adequately proved in any instance. 2. ROLES OF ENZYME SYNTHESIS AND DEGRADATION

The possibility that different forms of acetyl-CoA carboxylase might account for the dramatically different levels of enzyme activity in rat liver extracts following fasting and fat-free feeding have been considered by Majerus and Kilburn (96). Antibodies were prepared to avian liver carboxylase and shown to cross react with the rat liver carboxylase which was to be tested. In preliminary experiments it was shown that the antibody precipitated enzyme in the protomeric and the polymeric (citrate-treated) forms and also enzyme that had been treated with palmityl-CoA (dis­ aggregated, protomeric). Thus the antibody precipitated all forms of the enzyme that might be present in the different physiological conditions. Immunological analysis was carried out on acetyl-CoA carboxylase in liver extracts of rats in four conditions : high-fat fed for 1 week, fasted for 48 hours, Purina rat chow fed, and fat-free fed to 72 hours. These extracts

3.0

2.0

3 Lü

Έ M LU

IO

" 0 5 ~ " 1.0

2.0

3.0

4.0 ~

ENZYME ADDED ImU)

FIG. 5. Immunological analysis of acetyl-CoA carboxylase in rat liver extracts. The 105,000 g liver supernatant solutions pooled from three rats fed different diets were used. Aliquote of each of these preparations were assayed by adding 0.5 μ\ of rabbit antiserum to the initial incubation reactions. After prior incubation, these mixtures were centrifuged at 50,000 g for 10 minutes, and the acetyl-CoA carboxylase activity in the supernatant solution was measured. □ — □ , high-fat (45% fat) fed for 1 week; ■ — ■ , fasted 48 hours; O—O, Purina rat chow fed; # — # , fat-free fed 72 hours. From Majerus and Kilburn (96).

138

P. ROY VAGELOS

varied 25-fold in enzyme-specific activity. As seen in Fig. 5 when increasing amounts of each preparation were assayed in the presence of 0.5 μΐ of antiserum, and the acetyl-CoA carboxylase was measured in the super­ natant solutions after centrifugation, no activity was found in any of the preparations until 0.8-0.9 milliunit (mU) of enzyme was added. This indicates that all preparations had the same equivalence point : there was a constant amount of immunologically precipitable enzyme per unit of enzyme activity. This experiment indicates that the differences in specific activity of acetyl-CoA carboxylase seen upon dietary manipulation are due to different amounts of enzyme in the livers of these animals. A recent report by Nakanishi and Numa (106) has confirmed these findings and has shown that the decrease in acetyl-CoA carboxylase activity observed in livers of diabetic rats is also due to a decreased quantity of the carboxylase protein in this condition. Thus the differences in carboxylase activity in rats fed different diets and in diabetic rats can be explained on the basis of differences in amounts of enzyme, and inhibition of enzyme by increased liver concentrations of palmityl-CoA does not play a role in this phe­ nomenon as measured in vitro. However these results do not exclude the possibility that regulation also occurs by changes in the catalytic efficiency per enzyme molecule due to various metabolites (tricarboxylic acids and palmityl-CoA) in vivo. In order to understand the role of synthesis and degradation in regulating the amount of acetyl-CoA carboxylase in rat liver during dietary manipula­ tions Majerus and Kilburn (96) have utilized the techniques of Schimke (133). The rates of enzyme synthesis were determined by quantitative precipitation of the enzyme by antibody after pulse labeling with leucine-3H. The rate of enzyme synthesis was increased 5- to 10-fold after fat-free feeding of previously fasted rats. The rates of degradation of the enzyme, expressed as half-life, were approximately 48 hours in rats fed a fat-free or 12% fat diet and 18 hours in fasted rats. Since the rate of enzyme synthesis was diminished in rats fed a 12% fat diet while the rate of degradation was similar to the fat-free animal, it is apparent that independent factors regulate the rates of acetyl-CoA carboxylase synthesis and degradation in rat liver. These results have been confirmed and extended to the diabetic state by Nakanishi and Numa (106), who found that the rate of enzyme synthesis was decreased in the diabetic rat. The rate of degradation of the enzyme was the same in normal, refed, and diabetic states, but it was faster in fasted rats. Thus the increase or decrease in enzyme quantity in rats on a fat-free or 12% fat diet or in diabetic rats is due primarily to increased or decreased rate of enzyme synthesis. The decrease in amounts of enzyme in fasted rats, which are not in a steady state, is due to both decreased enzyme synthesis and increased enzyme degradation.

REGULATION OF FATTY ACID BIOSYNTHESIS

139

The involvement of protein synthesis in the rise in fatty acid synthesis seen upon refeeding fasted rats was suggested by studies of Allman, Hubbard, and Gibson (10), who showed that administration of actinomycin D before refeeding prevented the rise in fatty acid synthesis. Although the increase in acetyl-CoA carboxylase synthesis upon refeeding has now been demonstrated (96, 106), the inducer of enzyme synthesis is unknown. In this connection it should be noted that Allman et al. (10) demonstrated that liver linoleate levels parallel the changes in fatty acid synthesis induced by diet. During fat-free feeding, linoleate levels in liver fall rapidly. Administration of small amounts of linoleate in the fat-free diet can prevent the full induction of fatty acid synthesis while the feeding of other lipids has no effect. The mechanism of this linoleate effect is not understood. Majerus et al. (95) have shown in well differentiated rat hepatomas that neither acetyl-CoA carboxylase nor fatty acid synthetase is subject to the changes in enzyme activity normally observed in the host liver during dietary manipulations. Acetyl-CoA carboxylase was purified from both the hepatoma and the host liver and compared. Since the enzyme from both sources appeared similar with respect to activation by citrate and inhibition by palmityl-CoA, the authors concluded that the enzyme in hepatoma and liver is the same; the reason that the hepatoma enzyme does not respond to dietary manipulations is that these conditions do not affect the rate of enzyme synthesis and degradation in the hepatoma whereas they do affect these processes in liver. Preliminary experiments have shown that there is no change in the rate of acetyl-CoA carboxylase synthesis in hepatoma after fat-free feeding while there is a marked increase in synthesis of the enzyme in the host liver (96). Dakshinamurti and Desjardins have reported experiments which indicate that acetyl-CoA carboxylase responds to biotin deficiency differently in rat liver and adipose tissue (30, 31). They found that the adipose tissue enzyme activity was decreased 6-fold during biotin deprivation whereas the liver enzyme activity was only 50% decreased. Examination of the lipids of these two organs during biotin deprivation indicated that the lipid content of adipose tissue was markedly reduced while the liver lipids showed minimal changes, reflecting the changes in acetyl-CoA carboxylase activity. These observations have been followed up by Jacobs, Kilburn, and Majerus (63), who have demonstrated that the reduction in acetyl-CoA carboxylase activity in adipose tissue during biotin deficiency is due to conversion of the holoenzyme to apoenzyme. Thus acetyl-CoA carboxylase activity in adipose tissue is regulated not only by changes in rates of synthesis and degradation of the protein, but also by the relative concen­ trations of apoenzyme and holoenzyme which are determined by the availability of biotin.

140

P. ROY VAGELOS

3. REPRESSION

Acetyl-CoA carboxylase of Lactobacillus plantarum is regulated by repres­ sion (13, III). Long-chain unsaturated fatty acids, such as oleate or eisvaccenate at concentrations of 50 Mg/ml, cause severe repression of the carboxylase. Saturated fatty acids, such as palmitate or stéarate, are without effect. Since long-chain fatty acids are the end products of the fatty acid biosynthetic pathway, it is not surprising that these compounds repress the first committed enzyme of the pathway. It will be noted below that the enzymes of the fatty acid synthetase are also subject to repression by fatty acids in this organism. It is perhaps more surprising that the fatty acid biosynthetic enzymes in other microorganisms such as Escherichia coli do not appear to be sensitive to repression by exogenous long-chain fatty acids. Because the L. plantarum carboxylase is sensitive to repression by long-chain fatty acids, the level of this enzyme can be regulated by the concentration of biotin in the growth medium. When the organism is grown under conditions of biotin deficiency, the acetyl-CoA carboxylase holoenzyme activity is very low, and this leads to decreased synthesis of longchain fatty acids. The synthesis of the apoenzyme is derepressed in these cells, and its concentration rises to a level approximately 3-fold higher than that found in cells grown on sufficient levels of biotin. On the other hand, culture of the organism with excess biotin in the growth medium results in the rapid synthesis of long-chain unsaturated fatty acids which severely depress the level of enzyme synthesis. The effects of biotin on enzyme synthesis appear to be secondary since the long-chain unsaturated fatty acids (or their thioester derivatives) are the true repressore in this system. 4. PHARMACOLOGICAL INHIBITORS

A number of hypolipidemic agents, namely 2-methyl-2-[p-l,2,3,4tetrahydro-1-naphthyl) phenoxy]propionate, ethyl 2- (p-chlorophenyl) -2methylpropionate, and 2-methyl-2-[(p-chlorophenyl) phenoxy]propionate have been found by Maragoudakis to inhibit acetyl-CoA carboxylase (98-100). All three of these compounds have been reported to be effective drugs in reducing the plasma levels of cholesterol-rich and triglyceride-rich lipoproteins. Kinetic analyses of all three agents, using avian or rat liver acetyl-CoA carboxylases have indicated that inhibition is competitive for both the substrate acetyl-CoA and the activator isocitrate and noncompetitive for ATP and HC0 3 ~. It is of interest that the isocitratedependent aggregation of the carboxylase is reversed by these agents. These drugs do not inhibit fatty acid synthetase.

R E G U L A T I O N O F FATTY ACID

BIOSYNTHESIS

141

III. Fatty Acid Synthetase A. General Biosynthetic Scheme The enzyme system which catalyzes the synthesis of saturated long-chain fatty acids from malonyl-CoA is called the fatty acid synthetase (80, 85, 97), and the requirements of this reaction include acetyl-CoA and NADPH, as indicated in Eq. (13) : CH3CO—S—CoA + 7HOOCCH2CO—S—CoA + 14NADPH + 14H+ -* CH3CH2(CH2CH2)6CH2COOH + 7C0 2 + 14NADP+ + 8C0A—SH + 6H 2 0

(13)

This enzyme system was discovered by Lynen and his associates in yeast, where it is present as a multienzyme complex (80, 85, 97). Com­ parable multienzyme complexes were later isolated from avian liver (59), rat liver (22), rat mammary gland (HI), and Mycobacterium phlei (19). The absence of free intermediates, susceptibility of the enzyme complex to inhibition by sulfhydryl poisons, and the identification of protein-bound acetoacetate led Lynen to propose a mechanism of fatty acid biosynthesis in which all of the intermediates are acyl-protein derivatives. The proposed intermediates were based upon studies with model compounds (80). The nature of the protein-bound intermediates as well as the details of the intermediate reactions were elucidated in studies of bacterial systems, initially Clostridium kluyveri and then Escherichia coli (97, 159, 163). In contrast to the tightly associated multienzyme complexes listed above, the fatty acid synthetase of these microorganisms was found dissociated when the cells were disrupted, and the individual proteins of the synthetase have been isolated and studied. Although these bacterial systems have facilitated the understanding of fatty acid biosynthesis, they have posed an interesting biological problem: Are the bacterial fatty acid synthetase proteins associated in vivo in a typical multienzyme complex? All attempts to isolate a complex from E. coli have failed thus far. Preliminary studies, however, have shown that acyl carrier protein (ACP), a central component of the fatty acid synthetase, is localized on or near the inside surface of the plasma membrane in E. coli. The fact that ACP is not randomly distributed in the cell suggests a degree of organization which was not appreciated before, and this finding suggests that the E. coli fatty acid synthetase may be organized in vivo, perhaps in a typical multienzyme complex which includes ACP (165). Studies of the individual enzymes of the E. coli fatty acid synthetase have indicated the following reactions which explain the biosynthetic sequence from malonyl-CoA to the major saturated product, palmitate

142

P. ROY VAGELOS

{92, 97, 163, 173) : CH3CO—S—CoA + HS—ACP ^ CH3CO—S—ACP + CoA—SH

( 14)

CH3CO—S—ACP + HS—Econd. ^± CH3CO—S—Econd. + HS—ACP

( 15)

HOOCCH2CO—S—CoA + HS—ACP ^± HOOCCHoCO—S—ACP + CoA—SH

( 16)

HOOCCH2CO—S—ACP + CH3CO—S—Econd. ^ C0 2 + HS—Econd. + CH3COCH2CO—S—ACP

(17)

CH3COCH2CO—S—ACP + NADPH + H+ ;=± NADP+ + CH3CHOHCH2CO—S—ACP

( 18)

CH3CHOHCH2CO—S—ACP ^± CH3CH=CHCO—S—ACP + H 2 0

( 19)

CH3CH=CHCO—S—ACP + NADPH + H+ ^± NADP+ + CH3CH2CH2CO—S—ACP

(20)

In the initial reaction (Eq. 14) an acetyl group is transferred by acetylCoA-ACP transacylase from the sulfhydryl group of CoA to the single sulfhydryl group of ACP to form acetyl-ACP. The acetyl group is then transferred to a sulfhydryl group of the condensing enzyme (Econd.) to form an acetyl-enzyme intermediate and liberate ACP (Eq. 15). MalonylCoA-ACP transacylase catalyzes the transfer of a malonyl group from CoA to ACP in Eq. 16. This is followed by the condensation reaction (Eq. 17), which takes place between malonyl-ACP and acetyl enzyme to produce acetoacetyl-ACP, C0 2 , and the free condensing enzyme. Acetoacetyl-ACP is then reduced by NADPH to form specifically D-( — )-ß-hydroxybutyrylACP (Eq. 18) ; the latter is dehydrated to form the trans unsaturated thioester, crotonyl-ACP (Eq. 19) ; and crotonyl-ACP is reduced by NADPH to form the saturated thioester, butyryl-ACP (Eq. 20). In the normal biosynthetic sequence butyryl-ACP reacts with the condensing enzyme to form butyryl enzyme (Eq. 15) thereby liberating ACP which can accept a malonyl group and thus initiate another elongation, reduction, dehydration, reduction sequence. After appropriate repetitions of this series of reactions, the normal product, palmityl-ACP is formed. Small amounts of myristate and stéarate are also produced, however the major saturated fatty acid produced in vivo and in vitro contains 16 carbon atoms. The in vitro products of the E. coli fatty acid synthetase are free fatty acids, and this is probably due to thioester hydrolysis catalyzed by a specific palmityl thioesterase which has been characterized by Barnes and Wakil {12), In vivo palmityl-ACP and other long-chain acyl-ACP's produced by

143

REGULATION OF FATTY ACID BIOSYNTHESIS

the synthetase probably react directly with glycerol 3-phosphate and a specific membranous acyltransferase to form lysophosphatidic acid, the first intermediate in the pathway of phospholipid biosynthesis (4, 43, 44, 164)Localization of ACP on the inner surface of the membrane (165) provides a convenient arrangement for transfer of the newly produced long-chain acyl groups into the phospholipids which are major components of the cell membrane. B. Acyl Carrier Protein The central role that ACP plays in fatty acid biosynthesis is illustrated in Eqs. (14-20). The discovery, isolation, and demonstration of the im­ portant structural and functional properties of E. coli ACP have been recently reviewed (97, 163). The amino acid compositions of ACP's isolated from E. coli, C. butyricum, Arthrobacter, avocado, and spinach are listed in Table VI. A preliminary composition has also been reported for TABLE VI AMINO ACID COMPOSITION OF ACP

DERIVED FROM DIFFERENT SOURCES

Amino acid residues per molecule of protein

Amino acid Cysteic acid Lysine Histidine Arginine Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine /3-Alanine Taurine α

Vanaman et al. (166). Ailhaud et al. (6). e Simoni et al. (139).

b

coli0· 0 4 1 1 9 6 3 18 1 4 7 7 1 7 5 1 2 1 1

butyricum0 Arthrobacter0 0 4 2 0 13 1 3 14 1 1 6 7 3.58 8 7 1 3 1 1

0.08 5.1 1.0 1.2 13.9 1.8 5.3 10.0 0.8 5.0 12.0 6.0 0.82 5.9 5.8 0.13 2.8 0.96 0.87

Avocado c

Spinache

1.1 10.1 0.93 1.1 12.0 6.7 9.6 21.6 3.1 7.1 11.0 10.0 0.89 5.0 9.0 0.85 2.8 0.95 0.92

0.15 8.8 1.0 0.08 12.0 5.7 4.3 16.2 1.8 4.2 9.0 7.0 0.92 5.0 7.0 0.08 2.1 0.93 0.89

144

P . ROY VAGELOS

ACP isolated from yeast (179). The most significant feature of all ACP's is the prosthetic group, 4 / -phosphopantetheine, initially discovered in E. coli ACP (94-). This is represented by ß-alanine and taurine (the oxida­ tion product of 2-mercaptoethylanine) seen in the amino acid compositions of Table VI. It is the sulfhydryl group of 4'-phosphopantetheine with which the acyl groups are linked as thioesters in the reactions of fatty acid synthesis. The prosthetic group is covalently linked as a phosphodiester with a hydroxyl group of a serine residue of the peptide chain (93, 127). The complete amino acid sequence of E. coli ACP has been recently elucidated (166,167). It is apparent in Fig. 6 that the prosthetic group is on serine 36, approximately the middle of the 77 amino acid peptide. Peptide modification studies have begun to probe the aspects of ACP structure that are critical for function. Acetylation of the four lysine residues and the amino group of the terminal serine residue has no effect on the ability of ACP to function in fatty acid synthesis (91). Carboxypeptidase A treat­ ment of ACP yields peptide 1 through 74 lacking the single histidine of the molecule. This peptide (Table VII) is fully active in the malonyl-CoA-C02 exchange reaction, which is based upon the first three enzymes of fatty acid synthesis (90, 125). Trypsin treatment of ACP yields peptide 19 through 61, which is completely inactive in the CO2 exchange reaction (90). Cyanogen bromide treatment of ACP yields peptide 1 through 44 which is also inactive. Trypsin treatment of fully acetylated ACP yields peptide 7 through 77 since cleavage occurs only at the single arginine residue. This peptide, which lacks the amino terminal hexapeptide is completely inactive in the exchange reaction. Thus it is obvious that the peptide structure is very important for the activity of ACP with the enzymes of fatty acid biosynthesis. To further the understanding of the critical features of the structure of / 10 NH2-Ser-Thr-Ile-Glu-Glu-Arg-Val-Lys-Lys-Ile-Ile-Gly-Glu20 Gin -Leu-Gly-Val-Lys-Gln-Glu-Glu-Val

30

-Thr -Asp -Asn-Ala-Ser-

*

40

Phe - i/o I- Glu -Asp -L eu - Gly-Ala -Asp -Ser-Leu -Asp-Thr-Val-Glu 50 Leu-Val-Met-Ala-Leu-Glu-Glu-Glu-Phe-Asp-Thr-Glu-Ile-Pro60 Asp-Glu-Glu-Ala-Glu-Lys-lle-Thr-Thr-Vol-Gln-Ala-Ala-Ile70 77 Asp-Tyr-Ile-Asn-Gly-His-Gln-Alo-COOH

FIG. 6. The complete amino acid sequence of ACP from Escherichia coli. The asterisk indicates the position of the prosthetic group, 4'-phosphopantetheine. From Vanaman et al. (167).

145

REGULATION OF FATTY ACID BIOSYNTHESIS

TABLE VII PEPTIDES OF ACP

Activity

Peptide ACP (1 -» 77) 1 —* 74 19 —> 61 1 -► 44 7 —> 77

Method of preparation — Carboxypeptidase A Trypsin CNBr Trypsin treatment of acetylated ACP

Holo in malonyl-CoA-C02 exchange reaction Fully active Fully active Inactive Inactive Inactive

Apopeptide in Holo-ACP synthetase reaction Fully active Fully active Inactive Inactive

ACP, apo-ACP peptide 1 through 74 has been synthesized by the solid phase method of Merrifield (104) · When apo-ACP was converted enzymatically (see below) to active holo-ACP, the synthetic product was indistinguishable from authentic holo-ACP 1 through 74 on the basis of ion exchange chromatography, gel filtration on Sephadex G-50, and precipita­ tion by an antibody prepared against native ACP (54). Thus the stage is set for synthesizing apo-ACP peptides with appropriate substitutions. Studies of these peptides should define which amino acids are important for the function of ACP in fatty acid synthesis since all the enzymes of the E. coli fatty acid synthetase are available for testing the synthetic peptides. 1. SYNTHESIS AND TURNOVER

The synthesis of holo-ACP from apo-ACP and CoA is catalyzed by the enzyme, holo-ACP synthetase according to Eq. (21) : Mg2

CoA + apo-ACP

+

> holo-ACP + 3', 5'-adenosine diphosphate

(21)

This enzyme, which has been purified from extracts of E. coli (36, 125), has rigid structural requirements for both CoA and the apoprotein. Neither dephospho-CoA nor oxidized CoA can substitute for reduced CoA as 4'-phosphopantethéine donor with this enzyme. A number of apopeptides of ACP have been tested and, as indicated in Table VII, only apopeptide 1 through 74 is active. The same peptides of ACP which are active in the malonyl-CoA-C02 exchange reaction are also active in the holo-ACP synthetase reaction. Another E. coli enzyme, ACP hydrolase (162), catalyzes the hydrolysis of ACP to yield the prosthetic group and the apoprotein according to Eq. (22) : + Mn2

Holo-ACP

* apo-ACP + 4'-phosphopantetheine

(22)

146

P. ROY VAGELOS

Studies of this enzyme indicate that it is a highly specific phosphodiesterase ; although it catalyzes cleavage of 4'-phosphopantetheine from E. coli acetyl-ACP and ACP of C. butyricum, it is completely inactive with trypsin peptide 19 through 61 of E. coli ACP. The in vivo functioning of both holo-ACP synthetase and ACP hydrolase have been demonstrated by both pulse and pulse-chase experiments with labeled pantothenate in a pantothenate auxotroph of E. coli (124)- These studies established that CoA is the immediate precursor of ACP, that the 4'-phosphopantetheine prosthetic group turns over rapidly, and that the rate of ACP turnover is four times the rate of growth of the ACP pool. This striking turnover of the prosthetic group accomplishes no obvious purpose for the cell. However, similar turnover has recently been reported in studies of 4 / -phosphopantetheine of rat liver fatty acid synthetase (158). Further studies in E. coli have indicated that ACP is synthesized as a constant fraction of cell protein; thus it is produced constitutively (35). C. Enzymes of Fatty Acid Synthetase of Escherichia coli 1. TRANSACYLASES

Acetyl-CoA-ACP transacylase catalyzes the reaction of Eq. (14) (6, 181). Although longer-chain fatty acyl thioesters up to C8 can replace acetyl-CoA, the enzyme is relatively specific for the acetyl moiety since reaction rates with other chain lengths are much lower. Malonyl-CoA is inactive with this enzyme. Pantetheine can replace ACP, and acetylpantetheine readily substitutes for acetyl-CoA. The enzyme is severely inhibited by both iV-ethylmaleimide and iodoacetamide, and this inhibition is prevented by prior incubation with acetyl-CoA (181). Incubation of the enzyme with acetyl-l4C-CoA leads to the formation of acetyl- u C enzyme, which is separated from the reaction mixture by filtration through Sephadex G-25. The acetyl-l4C enzyme was able to transfer the acetyl-l4C group to either CoA or ACP. Although the nature of the acetyl- l4 C enzyme was not further characterized, Williamson and Wakil (181) have proposed that the enzyme intermediate is a thioester which functions as follows : Acetyl-S—CoA + HS-enzyme ^ acetyl-S-enzyme + CoA—S H

(23)

Acetyl-S-enzyme + HS—ACP ^ acetyl-S—ACP + HS-enzyme

(24)

The sum of these two reactions is the reaction of Eq. (14). Malonyl-CoA-ACP transacylase catalyzes the analogous transfer of a malonyl group (Eq. 16) (6, 181). This enzyme is very specific for the malonyl group since there is no activity with acetyl-CoA as a substrate. Pantetheine can substitute efficiently for ACP in this reaction, and malonyl pantetheine substitutes for malonyl-CoA. This transacylase is a sulfhydryl

REGULATION OF FATTY ACID BIOSYNTHESIS

147

FIG. 7. Phase contrast micrograph of single crystal of ß-ketoacyl-ACP synthetase.

enzyme as it is stimulated by mercaptans and inhibited by maleimide and iodoacetamide. Incubation of the enzyme with 14 C-CoA leads to the formation of malonyl-14C enzyme which matically active {130), This acyl enzyme intermediate has not characterized.

iV-ethylmalonylis enzyyet been

2. CONDENSING ENZYME

The condensation reaction of fatty acid synthesis is catalyzed by ß-ketoacyl-ACP synthetase (Eqs. 15 and 17) (7, 154) which has been recently purified to homogeneity and crystallized {51, 126), Crystals of the enzyme (Fig. 7) are hexagonal bipyramids 0.18 mm in the longest dimension.

148

P . ROY VAGELOS

The molecular weight, determined from equilibrium centrifugation studies, is 66,000. Guanidine hydrochloride and sodium dodecyl sulfate cause dissociation of the enzyme into inactive subunits of approximately 34,000 daltons. This enzyme, like those above, is a sulfhydryl protein which is inhibited by alkylating agents. When the enzyme is incubated with iodoacetamide-14C under conditions that lead to 9 3 % inhibition, 100% of the radioactivity can be recovered as 14C-carboxymethylcysteine from an acid hydrolyzate of the alkylated enzyme. Acetyl-ACP completely protects the enzyme against alkylation. The enzyme reacts with acetyl- 3 H-ACP to form an acetyl-3H enzyme intermediate, and the properties of this inter­ mediate suggest that the acetyl group is bound to a cysteine residue as an acyl thioester (51 ). AcetyPH enzyme is active in transferring the acetyPH group either to ACP to form acetyl- 3 H-ACP (Eq. 15) or to malonyl-ACP to form acetoacetyl- 3 H-ACP (Eq. 17). It is therefore apparent that the first three enzymes of fatty acid biosynthesis, the two transacylases and the condensing enzyme, are sulfhydryl enzymes which catalyze the transfer of acyl groups. a. Enzyme Specificity in Regulation of Fatty Acid Chain Length. The pre­ dominating saturated fatty acid in E. coli is palmitate whereas the pre­ dominating unsaturated fatty acids are palmitoleate and m-vaccenate (70, 77). The fatty acid synthetase of this organism synthesizes both saturated and unsaturated fatty acids in vitro and, as shown by Bloch (15) and his co-workers, the critical reaction in the biosynthetic pathway that leads to unsaturated fatty acids is catalyzed by ß-hydroxydecanoyl thioester dehydrase. Figure 8 demonstrates the fact that ß-hydroxydecanoyl-ACP

ACETYL-COA + MALONYL-COA

CH3(CH2)5CH2CH0HCH2C0-S-ACP

/ H H CH3(CH2)5C=CCH2C0-S-ACP

C^iCHo^CHofkCO-S-ACP I

+ 3 Co H H CH3(CH2)5C=C(CH2)7C0-S-ACP PALMITOLEATE

I T

H H

MADPH t CH3(CH2)5CH2CH2CH2C0-S-ACP I

+

C2

PALMITATE

CH3(CH2)5C=C(CH2)9C0-S-ACP £1£-VACCENATE

FIG. 8. Fatty acid biosynthesis in Escherichia colt. Reaction A initiates the pathway to unsaturated acids and reaction B, the pathway to saturated acids.

149

REGULATION OF FATTY ACID BIOSYNTHESIS TABLE VIII ACTIVITIES OF /3-KETOACYL A C P

SYNTHETASE WITH VARIOUS

INTERMEDIATES OF F A T T Y ACID SYNTHESIS 0

Intermediate Acetyl-ACP Deeanoyl-ACP Dodecanoyl-ACP Tetradecanoyl-ACP Hexade canoy 1-ACP cis-3-Decenoyl-ACP czs-5-Dodecenoyl-ACP czs-9-Hexadecenoyl-ACP cis-11-Octadecenoyl-ACP

Km (μΜ) 0.52 0.33 0.27 0.28

—

0.71 0.20 0.37

—

'max (μΓΠθΙββ

product/min/mg) 2.8 2.8 0.97 0.31 NA& 1.9 1.7 0.37 NA

α Assays were carried out according to Greenspan et al. (53). Kinetic constants were determined from Lineweaver-Burk plots of the data obtained. 6 NA, no activity.

is the intermediate at the branch point between the pathways to saturated and unsaturated fatty acids. Dehydration of this compound in pathway A gives rise to m-3-decenoyl-ACP. This thioester presumably condenses with malonyl-ACP to initiate chain elongation of m-unsaturated acyl-ACP intermediates. Other hypothetical intermediates expected to undergo condensation with malonyl-ACP in this pathway include czs-5-dodecenoylACP, as-7-tetradecenoyl-ACP, and cis-9-hexadecenoyl-ACP (palmitoleate). On the other hand, pathway B is initiated by a dehydration which forms 2r There are 4 moles of FMN per mole of enzyme complex. ACP is the only protein which has been isolated from the complex and * Recent experiments suggest that the yeast synthetase contains 3.5-6.0 moles of 4'-phosphopantetheine per mole of complex (185).

152

P. ROY VAGELOS

identified. The presence of a protein containing covalently bound 4'-phosphopantetheine was initially suggested when one of the peptides derived from proteolytic digestion of the complex was found to contain equimolar quantities of 0-alanine, 2-mercaptoethylamine, and phosphate. A pantothenic acid-requiring strain of baker's yeast was grown in medium con­ taining pantothenate- 14 C, and the enzyme complex was isolated and shown to contain 3 moles of 4'-phosphopantetheine- 14 C. Release of the 4'-phosphopantetheine from the enzyme by treatment with alkali was consistent with behavior expected for serine-bound phosphodiesters (176). These studies, demonstrating the presence of protein-bound 4'-phosphopantetheine in a multienzyme complex, were similar to studies reported earlier with the fatty acid synthetase complex of rat adipose tissue (76). The yeast ACP was isolated after denaturation of pantetheine-14C fatty acid synthetase in guanidine hydrochloride (179). After alkylation of the labeled protein with iodoacetamide, it was isolated by Sephadex filtration in guanidine hydrochloride and preparative polyacrylamide gel electrophoresis in phenol-containing medium. The final step of purification gave rise to two pantetheine- l4 C containing proteins on gel electrophoresis; however, both of these had very similar amino acid compositions. The finding that two proteins appear to represent yeast ACP is not yet under­ stood. The molecular weight of yeast ACP is estimated to be 16,000 from the amino acid analyses. Unfortunately, since the isolation procedure in­ cluded alkylation of the protein, active ACP could not be derived by this method. It should be noted that yeast synthetase is the only multienzyme complex from which ACP has been isolated thus far. b. Function. The overall reaction catalyzed by yeast fatty acid synthetase is shown in Eq. 25 : CH3CO—S—CoA + rcHOOCCH2CO—S—CoA + 2nNADPH + 2nH+ -> CH3(CH2CH2)„CO—S—CoA + nC0 2 + nCoA—SH + 2rcNADP+ + wH20

(25)

The products of the reaction are CoA esters of palmitate and stéarate. As individual active enzymes have not been isolated, details of the reaction mechanism have been derived primarily from studies of model reactions catalyzed by the complex and studies of acyl complex intermediates. The scheme shown in Fig. 10 indicates the reaction mechanism postulated by Lynen. Two types of sulfhydryl groups are indicated on the scheme. One has been denoted the "central" thiol and that is indicated by bold type; the other is the "peripheral" thiol group (80, 85). In this sequence an acetyl group is transferred in the priming reaction to the "peripheral"

„

s

CHa.LCHa.CH^+KCO.Sx /Enzyme HS-"

^ *

^

CH3.[CH2.CH2]B.CH(OH).CH2.CO.S. ^ Enzyme+NADP+ HS

; = ±

.

HSX / E n z y m e + CH 3 · [CH 2 · CH 2 ],, + r CO- SCoA H8'

HS\ .Enzyme

CHa.tC^.CH^+i.CO.S - — ^ E n z y m e + NADP+ HS'

CHa.tCHî.CH^+i.CO.S^

(FMN) ""*">

CH3.[CH2.CH2]ll.CH:CH.CO.S. ^ Enzyme+ H 2 0 HS

,

FIG. 10. Mechanism of fatty acid synthesis in yeast. From Lynen {85),

CHa.tCHa.CHzk+i.CO.Sv ^ E n z y m e + HSCoA HS^

.

CHa.LCHz.CH^.CHiCH.CO.S. "' "" ^ E n z y m e + NADPH + H+ HS^

CH 3 .[CH 2 -CH 2 ]«.CH(OH).CH 2 .CO.S. ^Enzyme HS

Terminal reaction :

(6)

(5)

(4)

CH34CH2.CH2ln.CO.CH2.CO.Sv. ^ E n z y m e + N A D P H + H+ HS

CH3-[CH2.CH2]n.C(0).CH2.CO.S\ „ 0 / Enzyme + C 0 2 HS

(3)

^

*

I ^ CH2.CO.S\ / E n z y m e + HSCoA CH3-[CH2.CH2]n.CO.Sx

: Enzyme + HSCoA

CHi-CO-S. ^Enzyme CHa.LCHj.CHrf.-CO.S^

I

C02H

-,

HSx

HS\ ^Enzyme CH3.[CH2.CH2]„.CO.Sx

^

^

(2)

(1)

C0 2 H I CH 2 -CO-SCoA +

Chain-lengthening reactions:

HSv CH3.CO.SC0A + /Enzyme

Priming reaction:

^

§3

h-*

Φ

GO

a

2 § ^

Ξ S

"* >. O g

%

g ^ *j

O

S £

tì

154

P. ROY VAGELOS

thiol. In reaction 1 of the chain-lengthening sequence a malonyl group is transferred to the "central" thiol. Condensation takes place between the protein-bound malonyl and acetyl groups to form acetoacetyl protein, this product occupying the "central" thiol (reaction 2). Reactions 3, 4, and 5 represent a reduction, dehydration, and reduction of the acyl group on the "central" thiol. Reduction of the enoyl protein derivative by NADPH in reaction 5 requires FMN. In reaction 6 the acyl group is transferred from the "central" malonyl group so that it can accept another malonyl group in reaction 1. This sequence is repeated until long-chain saturated fatty acids (Ci6 or Ci8) are formed. At this point the palmityl or stearyl group is transferred from the "central" thiol to CoA to form palmityl-CoA or stearyl-CoA (terminal reaction). With the identification of the "peripheral" thiol as a cyst eine residue of the condensing enzyme (81, 83, 87, 89) and the "central" thiol as the 4 / -phosphopantetheine of ACP (97), the striking similarity between this scheme and the reactions delineated in E. coli (Eqs. 14-20) is obvious. One major difference is the requirement for FMN in the yeast system; the other is that the products are thioesters of CoA in this system. i. "Central" and "peripheral" thiols. Two kinds of sulfhydryl groups were delineated in the yeast system on the basis of reactivity with alkylating agents and specific substrates. The "peripheral" thiol reacts very rapidly with iodoacetamide and can be protected from alkylation by this reagent by prior reaction with acetyl-CoA. Incubation of enzyme complex with acetyl-14C-CoA gives rise to 14C-acetyl enzyme which, upon incubation with malonyl-CoA, is converted to acetoacetyl-14C enzyme. This "periph­ eral" thiol has been shown to be a cysteine group of the condensing enzyme, and there are three such "peripheral" thiols per mole of yeast synthetase (81, 83, 87, 89). The "central" thiol, on the other hand, is much less sensi­ tive to iodoacetamide, and it has been identified as the 4'-phosphopantetheine of ACP (85). Although the sulfhydryl groups of the condensing enzyme and ACP have been denoted "central" and "peripheral" on the basis of relative sensitivity to iodoacetamide, the isolated pure condensing enzyme and ACP of E. coli also demonstrate similar differences in sensi­ tivity to alkylating agents (7). Thus the terms "central" and "peripheral" cannot be taken as an indication of the thiol locations within the multienzyme complex. ii. Transacylases. Performic acid oxidizes thioesters to the corresponding sulfonic acids with release of the carboxylic acids, and this has been utilized as a tool to study the amino acid side chains of the complex which bind acetyl and malonyl groups. When acetyl-14C enzyme, isolated after acetyll4 C-CoA reacted with the synthetase, was treated with performic acid, only 50% of the labeled acetyl groups were released. Those which remained

R E G U L A T I O N O F FATTY ACID

BIOSYNTHESIS

155

bound to th.e protein thus represented nonsulfhydryl acceptor groups in the protein. Similar results were obtained with malonyl-14C enzyme which was prepared and treated similarly. Studies of peptic hydrolyzates of malonylU C enzyme have corroborated the presence of two types of binding groups since two classes of peptides were obtained. In the first the malonyl group was released by performic acid; therefore this was a thioester and it was shown to contain 4 / -phosphopantetheine. In the second the malonyl group was stable to performic acid, and amino acids contained in a purified malonyl pentapeptide included serine, histidine, glycine, alanine, and leucine (85, 89). An overlapping malonyl heptapeptide was also isolated. Both the acyl pentapeptide and heptapeptide lacked cysteine and the malonyl group was very sensitive to alkaline hydrolysis; therefore Schweizer et al. {135) have concluded that the malonyl group is bound to serine by an O-ester linkage which is unusually activated by a nearby amino acid, possibly histidine. This malonyl binding site is thought to be the active center of the malonyltransferase of the complex. Peptic peptides of acetyl-14C enzyme have also been analyzed, and three classes of peptides have been elucidated (89). The first contained exclusively acetyl thioester with cysteine as the acetyl carrier. Since acetyl- l4 C groups were not bound by this site when the "peripheral" thiols were blocked by pretreatment of the enzyme complex with iodoacetamide, this cysteine site was assigned to the condensing enzyme. The second contained acetyl-14C thioesterified to 4'-phosphopantetheine. The third class of acetyl-14C peptides was stable to performic acid. Although the chemical composition of this third class was not reported, these acetyl peptides, which are said to be different from the performic acid stable malonyl peptides, are presumably part of the acetyl transacylase of the complex. Thus the yeast complex is thought to contain a malonyl trans­ acylase and an acetyl transacylase in which acyl groups are bound to the enzymes through nonthioester linkage. To summarize, malonyl groups are transferred from malonyl-CoA by the malonyl transacylase to the 4'-phosphopantetheine of ACP. Acetyl groups are transferred from acetyl-CoA to the 4 / -phosphopantetheine of ACP by acetyl transacylase. The acetyl groups are further transferred to a cysteine group of the condensing enzyme. Although this series of acyl transfers is similar to that described in the E. coli system, evidence suggests that both transacylases in E. coli are sulfhydryl enzymes, whereas the yeast complex enzymes contain acyl carrying residues which are not thiols. These differences require further investigation in both systems. iii. Regulation of fatty acid chain length. The products of the yeast synthetase are CoA esters of palmitate and stéarate, and the synthesis of these thioesters is catalyzed by a fatty acid transferase of the complex.

156

P. ROY VAGELOS

This transferase, which normally catalyzes the transfer of palmitate or stéarate from the complex to CoA (Eq. 26) has been studied by measuring the formation by the enzyme of labeled palmityl-CoA upon incubation of unlabeled palmityl-CoA with labeled CoA as in Eqs. (26)-(28), (80, 87). Palmityl-S—CoA + enzyme ^ palmityl-enzyme + CoA—SH Palmityl-enzyme +

C—CoA—SH

,4

^± palmityl- C—S—CoA + enzyme Sum:

Palmityl-S—CoA + 14

(26^

14

(27)

14

C—CoA—SH

;=± palmityl- C—S—CoA + CoA—SH

(28)

Since under standard conditions the synthetase forms almost exclusively fatty acids with a chain length of 16 to 18 C-atoms, Lynen, Hopper-Kessel, and Eggerer (87) suggested that this specificity might reflect a similar specificity of the transferase enzyme. If with shorter chain length fatty acyl groups the transferase reaction were slow compared to the condensation reaction, and if it became faster than the condensation reaction for chain lengths of 16 and 18 C-atoms, then palmitate and stéarate would be preferentially released from the enzyme. However, assay of the fatty acyltransferase activity with fatty acids of different chain lengths indicated that all saturated fatty acids with chain lengths between 6 and 18 C-atoms were transferred at about equal rates. Thus the specificity of this enzyme does not determine the chain length of the thioesters formed by the synthe­ tase (134). Because this transferase is resistant to inhibition by 2V-ethylmaleimide and iodoacetamide, the authors suggest that nonthiol substrate binding sites of the fatty acid synthetase complex are involved in the active site of this transferase. Recently Sumper et al. (146) have proposed a new model to explain termination at the level of Ci6 and Ci8 fatty acids in yeast. This model is based upon the assumption that chain elongation will terminate when the relative rate of the transferase reaction exceeds the rate of the condensing reaction. Interaction of the growing alkan chain must effect primarily the condensation reaction, since, as noted above, the transferase is nonspecific with regard to chain lengths between 6 and 18 carbon atoms. Thus chain termination appears to be due to the specificity of the condensing enzyme in the yeast synthetase as it is in E. coli. 2. LIVER

Pigeon liver fatty acid synthetase was isolated as a homogeneous complex by Hsu, Wasson, and Porter (59). It has a molecular weight of 450,000 (185); thus it is much smaller than the yeast complex. The product of

R E G U L A T I O N O F FATTY ACID

BIOSYNTHESIS

157

this enzyme is primarily palmitate, although small quantities of Cu and Ci8 fatty acids are also synthesized. The purified complex contains no flavin (59), but it contains 1 mole of 4 / -phosphopantetheine per mole of synthetase (24, 62, 180), presumably the prosthetic group of ACP. Al­ though minor quantities of the acetyl and malonyl transacylase activities dissociate from the complex when the latter is treated with 0.5 M guanidine hydrochloride, complete dissociation of the complex into its enzymatically active subunits has not been achieved. The complex (14.5 S) readily under­ goes dissociation into subunits (9.6 S) which have a molecular weight half of that of the original complex. Such dissociation occurs upon "aging" (184) or treatment of the complex with carboxymethyl disulfide, potassium maleate, or palmityl-CoA (24), and it is associated with loss of fatty acid synthetase activity. Complex dissociated by "aging" or treatment with carboxymethyldisulfide reassociates upon treatment with dithiothreitol to form a partially active complex with the same molecular weight as the original complex. Dissociation of the complex into half-molecular weight subunits by dialysis against Tris-glycine buffer, pH 8.3, containing 1 m l 2-mercaptoethanol or 1 mM dithiothreitol, on the other hand, is completely reversible since dialysis against 0.2 M potassium phosphate, pH 7.0, containing 10 mM dithiothreitol causes complete reassociation with a return to the same specific activity for fatty acid synthesis as the native complex (72). Analysis of the half-molecular weight subunits for the catalytic functions of the native complex, based on studying reactions of model compounds, has indicated that acetyl and malonyl transacylase, 0-ketoacyl and enoyl reductase, ß-hydroxyacyl dehydrase, and palmityl-CoA deacylase are all active. The most significant difference between the native and dissociated complex is the inability of the latter to catalyze the malonyl C0A-CO2 exchange reaction, which is a measure of the condensation reaction of fatty acid synthesis. When the synthetase is treated with a mixture of phenol, acetic acid, and urea followed by electrophoresis on polyacrylamide gel, at least eight peptides are seen; however, all catalytic activities are irreversibly lost. Although dissociation under these conditions indicates that the subunits of the complex are bound together by noncovalent forces only, it should be pointed out that treatment with dénaturants such as urea, guanidine hydrochloride, or detergents do not dissociate this complex (185). Direct measurement of NH 2 -terminal amino acids of the complex indicates that it contains at least five different DNP-amino acids. Acyl binding sites of the pigeon liver synthetase have been investigated and found to be similar to the binding sites discovered earlier in the yeast synthetase. There are three chemically distinct covalent binding sites for acetyl and two for malonyl moieties in the complex (27, 28, 62, 65,108,116,

158

P . ROY VAGELOS

117, 120), One thiol site, 4'-phosphopantetheine, and a nonthiol site bind both acetyl and malonyl groups, while another thiol site, cysteine, binds only acetyl groups. The nonthiol site, presumably in the active sites of the acetyl and malonyl transacylase, was identified as the hydroxyl group of a serine residue. Although the data suggest that the acetyl and malonyl groups bind to a common nonthiol site, it was impossible to determine in those experiments whether they bind to the same serine residue or whether each acyl group has a specific nonthiol site {65, 120). The cysteine site presumably resides in the active site of the condensing enzyme of the complex {65, 116). These sites are directly comparable to those of the yeast synthetase, and the proposed mechanism of acyl transfers and fatty acid synthesis is the same in the two systems {65, 108, 117, 120). The terminal reaction in the pigeon liver system is different from that in yeast since the reaction products are free fatty acids rather than thioesters. The liver synthetase contains a palmityl thioesterase which catalyzes cleavage of the fatty acid from the 4'-phosphopantetheine to yield the free synthetase and the long-chain fatty acid {12). This thioesterase has no activity with C12 or Ci4 saturated CoA derivations, good activity with palmityl-CoA, and decreased activity with stearyl-CoA. Rat liver fatty acid synthetase is very similar to the avian liver complex {22). It has a molecular weight of 540,000 with 1 mole of 4'-phosphopantetheine per mole of enzyme and no flavin. The two systems are also similar in that the integrity of the complexes requires the presence of thiol com­ pounds, they are both inhibited by sulfhydryl-binding reagents, and both form palmitate as the major product. One difference between the two is that the rat liver enzyme dissociates much more readily into lower molecular weight species in phosphate buffers below 0.5 M. However, complete dissociation into the active component enzymes has not been achieved. 3. MAMMARY GLAND

Fatty acid synthetase has been isolated from lactating rat {142) and rabbit mammary gland {25). Molecular weight of the rat enzyme is 478,000 and that of the rabbit is 910,000. The rat enzyme contains one 4'-phosphopantetheine per mole, and it readily dissociates into approximately halfmolecular weight subunits. With respect to turnover number and products of the reaction, the rat mammary enzyme is very similar to the complexes purified from pigeon and rat liver {142). The fact that homogenates from lactating rat {2) and rabbit {142) mammary gland synthesize both short and long-chain fatty acids via the malonyl-CoA pathway suggests that mammary tissue has a unique mechanism for terminating chain elongation by the fatty acid synthetase at shorter chain lengths. This mechanism must be extraneous to the fatty acid synthetase since the purified mammary complexes synthesize primarily palmitate.

R E G U L A T I O N O F FATTY ACID

BIOSYNTHESIS

159

4. MYCOBACTERIUM PHLEI

Mycobacterium phlei is an interesting bacterium in that it apparently has two independent fatty acid synthetases (20). The first is a multienzyme complex which has a molecular weight of 1.7 X 106 and contains 4'-phosphopantetheine. As reported with other complexes, this system is dissociated and inactivated by exposure to buffers of low ionic strength. The complex synthesizes straight-chain, even-numbered acids from Cu to C26 with a biphasic chain-length distribution, maxima occurring at d s and C24. The purified synthetase requires for activity two external supplements: one of these can be replaced by F M N ; the other has been separated into three polysaccharide fractions. The polysaccharides contain 3-O-methylmannose, mannose, 6-O-methylglucose, and glucose. The polysaccharides exert their effect on the fatty acid synthetase by lowering the Km for acetyl-CoA about 50-fold (60). There is another fatty acid synthetase in M. phlei extracts wThich has an average molecular weight less than 250,000. This activity is completely dependent upon the addition of ACP from either M. phlei or E. coli. This synthetase is unique in that it uses only palmityl-CoA or stearyl-CoA, but not acetyl-CoA, for chain initiation. It is not yet clear whether this second system might be derived from dissociation of the multienzyme complex (20). E. Control Early studies suggested that regulation of fatty acid biosynthesis might be mediated entirely through effects on acetyl-CoA carboxylase since liver extracts were found to contain acetyl-CoA carboxylase levels that were less than one-tenth the levels of fatty acid synthetase (38,110). However, more recent experiments, in which optimal conditions were utilized for car­ boxylase assay, have indicated that liver extracts of mouse, rat, and laying hen contain nearly equal carboxylase and synthetase activities (26). In addition numerous reports have established that liver fatty acid synthetase activity is diminished when an animal is fasted (10, 23, 1^1) and that it rises to levels far above normal when fat-free diets are used in realimentation (10, 40). Enzyme activity is also decreased in livers of alloxan-diabetic rats (29, J^l ), and it is corrected by the in vivo administration of insulin. Thus investigators have sought to explain the changes in levels of enzyme activity in these conditions on the basis of allosteric phenomena or changes in enzyme concentrations. 1. ALLOSTERIC REGULATION

a. Fructose 1,6-Diphosphate. The observation that potassium phosphate or phosphorylated sugars markedly stimulate pigeon liver fatty acid

160

P. ROY VAGELOS

synthetase is of interest in light of the recognized relationship between carbohydrate metabolism and fatty acid synthesis {119, 172). Although fructose 1,6-diphosphate caused the greatest stimulation, at higher con­ centrations glucose 1-phosphate, glucose 6-phosphate, glycerol 3-phosphate, pyrophosphate, and orthophosphate wrere also stimulatory to some degree. Initial experiments suggested that fructose 1,6-diphosphate is an allosteric activator of the fatty acid synthetase and that its mode of action was the reduction of the apparent Km for acetyl-CoA (172). Later studies showed that the enzyme preparation used in the initial experiments was con­ taminated with malonyl-CoA decarboxylase, and the effects on the Km for acetyl-CoA were not observed with a preparation that lacked this contaminant. Kinetic studies with the highly purified synthetase showed that the enzyme is sensitive to inhibition by malonyl-CoA, one of the substrates of the enzyme. The inhibition is of the mixed type with respect to acetyl-CoA and is competitive with respect to NADPH. Malonyl-CoA increases the Km for NADPH 19-fold over a malonyl-CoA concentration range of 10-37.5 μΜ. Inhibition by malonyl-CoA is reversed by fructose 1,6-diphosphate, which causes a decrease in the Km for NADPH. Fructose 1,6-diphosphate does not markedly effect the Km values for either acetylCoA or malonyl-CoA. The kinetic patterns with malonyl-CoA inhibition suggest that inhibition by this compound is due to substrate binding at an allosteric site (119). The physiological significance of these observations is not yet clear since tissue concentrations of malonyl-CoA have not been reported. b. Feedback Inhibition. Palmityl-CoA and other long-chain acyl-CoA compounds inhibit fatty acid synthetase of pigeon liver (33, 122), rat liver (156, 157), rat brain (128), and yeast (79). Kinetic experiments with the yeast enzyme have indicated that all long-chain acyl-CoA thioesters tested were competitive inhibitors of the substrate, malonyl-CoA, and Lust and Lynen have proposed that these compounds regulate fatty acid biosynthesis via end product inhibition (79). However, as in the case of acetyl-CoA carboxylase inhibition by long-chain acyl-CoA compounds, the physiological importance of the in vitro observations has been ques­ tioned. A critical evaluation of the effect of palmityl-CoA on pigeon liver synthetase by Dorsey and Porter (33) has indicated that inhibition of the enzyme is dependent on the presence of a critical mixed micellar concentra­ tion of this thioester and, more importantly, on the molar ratio of the CoA ester to protein. The molar ratio requirement seems to eliminate the possibility of site-specific inhibition of this enzyme by palmityl-CoA. These authors conclude that palmityl-CoA inhibits this enzyme by virtue of its detergent nature, and their proposal is supported by their finding that sodium lauryl sulfate, another strong detergent, inhibits the enzyme in a similar way.

REGULATION OF FATTY ACID BIOSYNTHESIS

161

2. ENZYME SYNTHESIS AND DEGRADATION

Although there may be some doubt as to the physiological role of the effectors of fatty acid synthetase studied in vitro, there is no doubt that the concentration of this enzyme varies in different nutritional and hor­ monal conditions. Porter and his co-workers {21, 23, 29) have measured the fatty acid synthetase concentration by purification of the protein to near homogeneity from livers of rats which were in various conditions. They found lowered levels of fatty acid synthetase in livers of fasted, alloxan diabetic and portacaval-shunted rats. Insulin treatment of alloxan diabetic rats caused a return of the level of the synthetase toward the normal range. In addition they found by the same types of measurements that the rise to supranormal levels, which is observed on feeding fasting rats a fat-free diet, is due to adaptive enzyme synthesis. This was shown by demonstrating that leucine-14C is readily incorporated in vivo into the purified fatty acid synthetase during refeeding. Thus, as in the case of acetyl-CoA carboxylase, fatty acid synthetase concentrations vary markedly under different nutritional and hormonal conditions. In the case of the synthetase the specific changes in rates of synthesis and degradation of the enzyme have not been studied in these conditions. However, recent measurements of the rate of turnover of this enzyme in steady-state animals have demon­ strated a half-life in the range of 71-108 hours {158) ; thus similar measure­ ments under the abnormal conditions discussed above should establish the important mechanisms which alter the turnover rate and determine the cellular concentration of this enzyme. 3. REPRESSION

Studies of the regulation of fatty acid biosynthesis in bacteria have indicated distinct differences in different organisms. The presence of longchain saturated or unsaturated fatty acids in growth medium does not effect fatty acid synthesis by Escherichia coli {132, 137, 138). On the other hand, Henderson and McNeill showed that fatty acid synthesis decreased when Lactobacillus plantarum was grown in the presence of unsaturated or cyclopropane fatty acids {56, 57). These authors found that maximal control in this organism requires a double bond or cyclopropane ring at carbons 9 or 11 of the fatty acid. Although the early experiments suggested that the mechanism underlying the decrease in fatty acid synthesis was feedback inhibition, more recent studies showed that extracts from oleategrown cells did not catalyze fatty acid synthesis; therefore enzyme repres­ sion was suggested as the important factor {57). These findings have been confirmed by Weeks and Wakil {175), who found that both acetyl-CoA carboxylase and fatty acid synthetase activities were reduced in extracts of this organism grown in the presence of oleate. Several of the individual enzymes of the fatty acid synthetase were reduced by 5- to 6-fold under

162

P. ROY VAGELOS

these conditions, suggesting that the synthesis of these enzymes is under some form of coordinate control. REFERENCES

1. Abraham, S., Matthes, K. J., and Chaikoff, I. L., J. Biol. Chem. 235, 2551 (1960). 2. Abraham, S., Matthes, K. J., and Chaikoff, I. L., Biochim. Biophys. Ada 46, 197 (1961). 3. Ahmad, F., Jacobson, B. E., and Wood, H. G., J. Biol. Chem. 245, 6486 (1970). 4. Ailhaud, G. P., and Vagelos, P. R., / . Biol. Chem. 241, 3866 (1966). 5. Ailhaud, G. P., Vagelos, P. R., and Goldfine, H., / . Biol. Chem. 242, 4459 (1967). 6. Alberts, A. W., Majerus, P. W., Talamo, B., and Vagelos, P. R., Biochemistry 3, 1563 (1964). 7. Alberts, A. W., Majerus, P. W., and Vagelos, P. R., Biochemistry 4, 2265 (1965). 8. Alberts, A. W., Nervi, A. M., and Vagelos, P. R., Proc. Nat. Acad. Sci. U. S. 63, 1319 (1969). 9. Alberts, A. W., and Vagelos, P. R., Proc. Nat. Acad. Sci. U. S. 59, 561 (1968). 10. Allman, D. W., Hubbard, D. D., and Gibson, D. M., / . Lipid Res. 6, 63 (1965). 11. Ballard, F. J., and Hanson, R. W., / . Lipid Res. 8, 73 (1967). 12. Barnes, E. M., Jr., and Wakil, S. J., / . Biol. Chem. 243, 2955 (1968). 13. Birnbaum, J., Arch. Biochem. Biophys. 132, 436 (1969). 14. Birnbaum, J., J. Bacteriol. 104, 171 (1970). 15. Bloch, K., Accounts Chem. Res. 2, 193 (1969). 16. Bortz, W. M., and Lynen, F., Biochem. Z. 337, 505 (1963). 17. Bortz, W. M., and Lynen, F., Biochem. Z. 339, 77 (1963). 18. Brady, R. O., and Gurin, S., / . Biol. Chem. 199, 421 (1952). 19. Brindley, D. N., Matsumura, S., and Bloch, K , Nature (London) 224, 666 (1969). 20. Bucker, N. L. R., J. Amer. Chem. Soc. 75, 498 (1953). 21. Burton, D. N., Collins, J. M., Kennan, A. L., and Porter, J. W., J. Biol. Chem. 244, 4510 (1969). 22. Burton, D. N., Haavik, A. G., and Porter, J. W., Arch. Biochem. Biophys. 126, 141 (1968). 23. Butterworth, P. H. W., Guchhait, R. B., Baum, H., Olson, E. B., Margolis, S. A., and Porter, J. W., Arch. Biochem. Biophys. 116, 453 (1966). 24. Butterworth, P. H. W., Yang, P. C., Bock, R. M., and Porter, J. W., J. Biol. Chem. 242, 3508 (1967). 25. Carey, E. M., and Dils, R., Biochim. Biophys. Ada 210, 371 (1970). 26. Chang, H.-C, Seidman, I., Teebor, G., and Lane, M. D., Biochem. Biophys. Res. Commun. 28, 682 (1967). 27. Chesterton, C. J., Butterworth, P. H. W., Abramowitz, A. S., Jacob, E. J., and Porter, J. W., Arch. Biochem. Biophys. 124, 386 (1968). 28. Chesterton, C. J., Butterworth, P. H. W., and Porter, J. W., Arch. Biochem. Biophys. 126, 864 (1968). 29. Dahlen, J. V., Kennan, A. L., and Porter, J. W., Arch. Biochem. Biophys. 124, 51 (1968). 30. Dakshinamurti, K , and Desjardins, P. R., Can. J. Biochem. 46, 1261 (1968). 31. Dakshinamurti, K., and Desjardins, P. R., Biochim. Biophys. Ada 176, 221 (1969). 32. Dimroth, P., Guchhait, R. B., Stoll, E., and Lane, M. D., Proc. Nat. Acad. Sci. U. S. 67, 1353 (1970). 33. Dorsey, J. A., and Porter, J. W., J. Biol. Chem. 243, 3512 (1968). 34. Eakin, R. E., Snell, E. E., and Williams, R. J., / . Biol. Chem. 136, 801 (1940). 35. Elovson, J., and Vagelos, P. R., unpublished observation.

REGULATION OF FATTY ACID BIOSYNTHESIS

36. 37. 38. 39.

163

Elovson, J., and Vagelos, P. R,, J. Biol. Chem. 243, 3603 (1968). Flatt, J. P., and Ball, E. G., / . Biol. Chem. 239, 675 (1964). Ganguly, J., Biochim. Biophys. Acta 40, 110 (1960). Gerwin, B. I., Jacobson, B. E., and Wood* H. G., Proc. Nat. Acad. Sci. U. S. 64, 1315 (1969). 40. Gibson, D. M., Hicks, S. E., and Allmann, D. W., Advan. Enzyme Regul. 4, 239 (1966). 41. Gibson, D. M., and Hubbard, D. D., Biochem. Biophys. Res. Commun. 3, 531 (1960). 42. Gibson, D. M., Titchner, E. B., and Wakil, S. J., J. Amer. Chem. Soc. 80, 2908 (1958). 43. Goldfine, H., / . Biol, Chem. 241, 3864 (1966). 44· Goldfine, H., Ailhaud, G. P., and Vagelos, P. R., / . Biol. Chem. 242, 4466 (1967). 45. Goto, T., Ringelmann, E., Reidei, B., and Numa, S., Life Sci. 6, 785 (1967). 46. Green, N. M., Biochem. J., 89, 585 (1963). 47. Gregolin, C., Ryder, E., Kleinschmidt, A. K., Warner, R. C., and Lane, M. D., Proc. Nat. Acad. Sci. U. S. 66, 148 (1966). 48. Gregolin, C , Ryder, E., and Lane, M. D., / . Biol. Chem. 243, 4227 (1968). 49. Gregolin, C., Ryder, E., Warner, R. C., Kleinschmidt, A. K., Chang, H. C., and Lane, M. D., J. Biol. Chem. 243, 4236 (1968). 50. Gregolin, C , Ryder, E., Warner, R. C., Kleinschmidt, A. K., and Lane, M. D., Proc. Nat. Acad. Sci. U. S. 66, 1751 (1966). 51. Greenspan, M. D., Alberts, A. W., and Vagelos, P. R., / . Biol. Chem. 244, 6477 (1969). 52. Greenspan, M. D., Birge, C. H., Powell, G. L., Hancock, W. S., and Vagelos, P. R., Science 170, 1203 (1970). 53. Greenspan, M. D., Birge, C. H., and Vagelos, P. R., Fed. Proc. 28, 537 (1969). 54. Hancock, W. S., Prescott, D. J., Nulty, W. L., Weintraub, J., Vagelos, P. R., and Marshall, G. R,, J. Amer. Chem. Soc. 93, 1799 (1971). 55. Heinstein, P. F., and Stumpf, P. K , / . Biol. Chem. 244, 5374 (1969). 56. Henderson, T. O., and McNeill, J. J., Biochem. Biophys. Res. Commun. 25, 662 (1966). 57. Henderson, T. O., and McNeill, J. J., Bacteriol. Proc. p. 134 (1967). 58. Himes, R. H., Young, D. L., Ringelmann, E., and Lynen, F., Biochem. Z. 337, 48 (1963). 59. Hsu, R. Y., Wasson, G., and Porter, J. W., J. Biol, Chem. 240, 3736 (1965). 60. Ilton, M., Jevans, A. W., McCarty, E. D., Vance, D., White, H. B., Ill, and Bloch, K , Proc. Nat. Acad. Sci. U. S. 68, 87 (1971). 61. Izui, K., Yoshinaga, T., Morikawa, M., and Katsuki, H., Biochem. Biophys. Res. Commun. 40, 949 (1970). 62. Jacob, E. J., Butterworth, P. H., and Porter, J. W., Arch. Biochem. Biophys. 124, 392 (1968). 63. Jacobs, R., Kilburn, E., and Majerus, P. W., J. Biol. Chem. 246, 6462 (1970). 64. Jacobson, B. E., Gerwin, B. L, Ahmad, F., Waegell, P., and Wood, H. G., J. Biol. Chem. 246, 6471 (1970). 65. Joshi, V. C , Plate, C. A., and Wakil, S. J., J. Biol. Chem. 24,5, 2857 (1970). 66. Klein, H. P., J. Bacteriol. 73, 530 (1957). 67. Kleinschmidt, A. K , Moss, J., and Lane, M. D., Science 166, 1276 (1969). 68. Knappe, J., Ringelmann, E., and Lynen, F., Biochem. Z. 336, 168 (1961). 69. Knappe, J., Wenger, B., and Wiegand, U., Biochem. Z. 337, 232 (1963). 70. Knivett, V. A., and Cullen, J., Biochem. J. 103, 299 (1967).

164

P. ROY VAGELOS

71. Kornacker, M. S., and Lowenstein, J. M., Biochem. J. 94, 209 (1965). 72. Kumar, S., Dorsey, J. A., Muesing, R. A., and Porter, J. W., J. Biol. Chem. 245, 4732 (1970). 78. Lane, M. D., and Lynen, F., Proc. Nat. Acad. Sci. U. S. 49, 379 (1963). 74. Lane, M. D., Vitamins Hormones 28, 345 (1971). 75. Langdon, R. G., / . Biol Chem. 226, 615 (1957). 76. Larrabee, A. R., McDaniel, E. G., Bakerman, H. A., and Vagelos, P. R., Proc. Nat. Acad. Sci. U. S. 64, 267 (1965). 77. Lennarz, W. J., Light, R. J., and Bloch, K , Proc. Nat. Acad. Sci. U. S. 48, 840 (1962). 78. Leveille, G. A., Fed. Proc. 29, 1294 (1970). 79. Lust, G., and Lynen, F., Eur. J, Biochem. 7, 68 (1968). 80. Lynen, F., Fed. Proc. 20, 941 (1961). 81. Lynen, F., Proc. Robert A. Welch Found. Conf. Chem. Res., Houston, Texas, 1962 5, 293. 82. Lynen, F., Biochem. Z. 337, 48 (1963). 88. Lynen, F., in "New Perspectives in Biology" (M. Sela, ed.), p. 132. Elsevier, Amsterdam, 1964. 84. Lynen, F., Angew. Chem. 77, 929 (1965). 85. Lynen, F., Biochem. J. 102, 381 (1967). 86. Lynen, F., Progr. Biochem. Pharmacol. 3, (1967). 87. Lynen, F., Hopper-Kessel, L, and Eggerer, H., Biochem. Z. 340, 95 (1964). 88. Lynen, F., Knappe, J., Lorch, E., Jutting, G., Ringelmann, E., and Lachance, J. P., Biochem. Z. 335, 123 (1961). 89. Lynen, F., Oesterhelt, D., Schweizer, E., Willecke, K., in "Cellular Compartmentalization and Control of F a t t y Acid Metabolism" (F. C. Gran, ed.), p. 1. Academic Press, New York, 1968. 90. Majerus, P. W., / . Biol. Chem. 242, 2325 (1967). 91. Majerus, P. W., Science 159, 428 (1968). 92. Majerus, P. W., Alberts, A. W., and Vagelos, P. R., Proc. Nat. Acad. Sci. U. S. 51, 1231 (1964). 98. Majerus, P. W., Alberts, A. W., and Vagelos, P. R., / . Biol. Chem. 240, 4723 (1965). 94. Majerus, P. W., Alberts, A. W., and Vagelos, P. R., Proc. Nat. Acad. Sci. U. S. 53, 410 (1965). 95. Majerus, P. W., Jacobs, R., Smith, M. E., and Morris, H. P., / . Biol. Chem. 243, 3588 (1968). 96. Majerus, P. W., and Kilburn, E., / . Biol. Chem. 244, 6254 (1969). 97. Majerus, P. W., and Vagelos, P. R., Advan. Lipid Res. 5, 1 (1967). 98. Maragoudakis, M. E., J. Biol. Chem. 244, 5005 (1969). 99. Maragoudakis, M. E., Biochemistry 9, 413 (1970). 100. Maragoudakis, M. E., / . Biol. Chem. 245, 4136 (1970). 101. Martin, D. B., and Vagelos, P. R., / . Biol. Chem. 237, 1787 (1962). 102. Matsuhashi, M., Matsuhashi, S., Numa, S., and Lynen, F., Fed. Proc. 21,288 (1962). 103. Matsuhashi, M., Matsuhashi, S., and Lynen, F., Biochem. Z. 340, 263 (1964). 104. Merrifield, R. B., / . Amer. Chem. Soc. 85, 2149 (1963). 105. Miller, A. L., and Levy, H. R., / . Biol, Chem. 244, 2334 (1969). 106. Nakanishi, S., and Numa, S., Eur. J. Biochem. 16, 161 (1970). 107. Nervi, A. M., Alberts, A. W., and Vagelos, P . R., Arch. Biochem. Biophys. 143, 401 (1971). 108. Nixon, J. E., Phillips, G. T., Abramovitz, A. S., and Porter, J. W., Arch. Biochem. Biophys. 138, 372 (1970).

REGULATION OF FATTY ACID BIOSYNTHESIS 109. 110. 111. 112. 118. 114. 115. 116. 117. 118. 119. 120. 121. 122. 123. 124. 125. 126. 127. 128. 129. 130. 131. 132. 133. 134135. 136. 137. 138. 139. 140. 141. 142. US. 144. 145. 146.

165

Numa, S., Bortz, W. M., and Lynen, F., Advan. Enzyme Regul. 3, 407 (1965). Numa, S., Matsuhashi, M., and Lynen, F., Biochem. Z. 334, 203 (1961). Numa, S., Ringelmann, E., and Lynen, F., Biochem. Z. 340, 228 (1964). Numa, S., Ringelmann, E., and Lynen, F., Biochem. Z. 343, 243 (1965). Numa, S., Ringelmann, E., and Reidei, B., Biochem. Biophys. Res. Commun. 24, 750 (1966). Oesterhelt, D., Bauer, H., and Lynen, F., Proc. Nat. Acad. Sci. U. S. 63, 1377 (1969). Pande, S. V., Khan, R. P., and Venkitasubramanian, T. A., Biochim. Biophys. Ada 84, 239 (1964). Phillips, G. T., Nixon, J. E., Abramovitz, A. S., and Porter, J. W., Arch. Biochem. Biophys. 138, 357 (1970). Phillips, G. T., Nixon, J. E., Dorsey, J. A., Butterworth, P. H. W., Chesterton, C. J7., and Porter, J. W., Arch. Biochem. Biophys. 138, 380 (1970). Pilz, L, Herbst, M., Kratky, O., Oesterhelt, D., and Lynen, F., Eur. J. Biochem. 13, 55 (1970). Plate, C. A., Joshi, V. C , Sedgwick, B., and Wakil, S. J., / . Biol. Chem. 243, 5439 (1968). Plate, C. A., Joshi, V. C , and Wakil, S. J., J. Biol. Chem. 245, 2867 (1970). Popjak, G., and Tietz, A., Biochem. J. 56, 46 (1954). Porter, J. W., and Long, R. W., / . Biol. Chem. 233, 20 (1958). Porter, J. W., Wakil, S. J., Tietz, A., Jacob, M. I., and Gibson, D. M., Biochim. Biophys. Ada 25, 35 (1957). Powell, G. L., Elovson, J., and Vagelos, P . R., / . Biol. Chem. 244, 5616 (1969) Prescott, D. J., Elovson, J., and Vagelos, P. R., / . Biol. Chem. 244, 4517 (1969). Prescott, D. J., and Vagelos, P. R., J. Biol. Chem. 245, 5484 (1970). Pugh, E. L., and Wakil, S. J., / . Biol. Chem. 240, 4727 (1965). Robinson, J. D., Brady, R, O., and Bradley, R. M., / . Lipid Res. 4, 144 (1963). Rognstad, R., and Katz, J., Proc. Nat. Acad. Sci. U. S. 55, 1148 (1966). Ruch, F., and Vagelos, P. R., unpublished observation. Ryder, E., Gregolin, C., Chang, H. C , and Lane, M. D., Proc. Nat. Acad. Sci. U. S. 57, 1455 (1967). Schairer, H. V., and Overath, P., / . Mol. Biol. 44, 209 (1969). Schimke, R, T., Bull. Soc. Chim. Biol. 48, 1009 (1966). Schweizer, E., Lerch, I., Kroeplin-Rueff, L., and Lynen, F., Eur. J. Biochem. 15, 472 (1970). Schweizer, E., Piccinini, F., Duba, C , Günther, S., Ritter, E., and Lynen, F., Eur. J. Biochem. 15, 483 (1970). Scrutton, M. C , Keech, D. B., and Utter, M. F., J. Biol. Chem. 240, 574 (1965). Silbert, D. F., Ruch, F., and Vagelos, P . R., J. Bacteriol. 95, 1658 (1968). Silbert, D . F., and Vagelos, P. R., Proc. Nat. Acad. Sei. U. S. 58, 1579 (1967). Simoni, R. D., Criddle, R. S., and Stumpf, P . K , J. Biol. Chem. 242, 573 (1967). Siperstein, M. D., and Fagan, V. M., J. Clin. Invest. 37, 1185 (1958). Smith, S., and Abraham, S., / . Biol. Chem. 245, 3209 (1970). Smith, S., and Dils, R., Biochim. Biophys. Ada 116, 23 (1966). Srere, P. A., Biochim. Biophys. Ada 106, 445 (1965). Srere, P. A., and Bhaduri, A., Biochim. Biophys. Ada 59, 487 (1962). Stell, E., Ryder, E., Edwards, J. B., and Lane, M. D., Proc. Nat. Acad. Sci. U. S. 60, 986 (1968). Sumper, M., Oesterhelt, D., Riepertinger, C , and Lynen, F., Eur. J. Biochem. 10, 377 (1969).

166

P. ROY VAGELOS

147. Sumper, M., Riepertinger, C, and Lynen, F., FEBS Lett. 5, 45 (1969). 148. Takeda, Y. H., Inoue, H., Honjo, K., Tanioka, H., and Daikuhara, Y., Biochim. Biophys. Ada 136, 214 (1967). 149. Taketa, K , and Pogell, B. M., / . Biol Chem. 241, 720 (1966). 150. Tepperman, H. M., and Tepperman, J., Diabetes 7, 478 (1958). 151. Tepperman, H. M., and Tepperman, J., Amer. J. Physiol. 206, 357 (1964). 152. Tepperman, J., and Tepperman, H. M., Amer. J. Physiol. 193, 55 (1958). 153. Tepperman, J., and Tepperman, H. M., Fed. Proc. 29, 1284 (1970). 154. Toomey, R. F., and Wakil, S. J., / . Biol. Chem. 241, 1159 (1966). 155. Tubbs, P. K , Biochim. Biophys. Ada 70, 608 (1963). 156. Tubbs, P. K., and Garland, P. B., Biochem. J. 89, 25P (1963). 157. Tubbs, P. K , and Garland, P. B., Biochem. J. 93, 550 (1964). 158. Tweto, J., Liberati, M., and Larrabee, A. R., / . Biol. Chem., in press. 159. Vagelos, P. R., Ann. Rev. Biochem. 33, 139 (1964). 160. Vagelos, P. R., Alberts, A. W., and Martin, D. B., Biochem. Biophys. Res. Commun. 8, 4 (1962). 161. Vagelos, P. R., Alberts, A. W., and Martin, D. B., J. Biol. Chem. 238, 533 (1963). 162. Vagelos, P. R., and Larrabee, A. R., / . Biol. Chem. 242, 1776 (1967). 168. Vagelos, P. R., Majerus, P. W., Alberts, A. W., Larrabee, A. R., and Ailhaud, G. P., Fed. Proc. 26, 1485 (1966). 164. van den Bosch, H., and Vagelos, P. R., Biochim. Biophys. Ada 218, 233 (1970). 165. van den Bosch, H., Williamson, J. R., and Vagelos, P. R,, Nature {London) 228, 338 (1970). 166. Vanaman, T. C , Wakil, S. J., and Hill, R. L., J. Biol. Chem. 243, 6409 (1968). 167. Vanaman, T. C, Wakil, S. J., and Hill, R. L., / . Biol. Chem. 243, 6420 (1968). 168. Waite, M., and Wakil, S. J., / . Biol. Chem. 237, 2750 (1962). 169. Waite, M., and Wakil, S. J., J. Biol. Chem. 238, 77 (1963). 170. Wakil, S. J., J. Amer. Chem. Soc. 80, 6465 (1958). 171. Wakil, S. J., in "Lipid Metabolism" (S. J. Wakil, ed.), pp. 1-48. Academic Press, New York, 1970. 172. Wakil, S. J., Goldman, J. K , Williamson, I. P., and Toomey, R. E., Proc. Nat. Acad. Sci. U. S. 55, 880 (1966). 178. Wakil, S. J., Pugh, E. L., and Sauer, F., Proc. Nat. Acad. Sci. U. S. 52, 106 (1964). 174. Wakil, S. J., Titchner, E. B., and Gibson, D. M., Biochim. Biophys. Ada 29, 225 (1958). 175. Weeks, G., and Wakil, S. J., J. Biol, Chem. 246, 1913 (1970). 176. Wells, W. W., Schultz, J., and Lynen, F., Proc. Nat. Acad. Sci. U. S. 56, 633 (1966). 177. Wessman, G. E., and Werkman, C. H., Arch. Biochem. Biophys. 26, 214 (1950). 178. Wieland, O., and Weiss, L., Biochem. Biophys. Res. Commun. 13, 26 (1963). 179. Willecke, K., Ritter, E., and Lynen, F., Eur. J. Biochem. 8, 503 (1969). 180. Williamson, I. P., Goldman, J. K , and Wakil, S. J., Fed. Proc. 25, 340 (1966). 181. Williamson, I. P., and Wakil, S. J., J. Biol. Chem. 241, 2326 (1966). 182. Wise, E. M., Jr., and Ball, E. G., Proc. Nat. Acad. Sci. U. S. 52, 1255 (1964). 183. Wood, H. G., Lochmuller, H., Riepertinger, G., and Lynen, F., Biochem. Z. 337, 247 (1963). 184. Yang, P. C., Bock, R. M., Hsu, R. Y., and Porter, J. W., Biochim. Biophys. Ada 110, 608 (1965). 185. Yang, P. C., Butterworth, P. H. W., Bock, R. M., and Porter, J. W., J. Biol. Chem. 242, 3501 (1967). 186. Young, J. W., Shrago, E., and Lardy, H. A., Biochemistry 3, 1687 (1964).

Kinetic Analysis of Allosteric Enzymes KASPER

KIRSCHNER

Department of Biochemistry Stanford University Stanford, California I. Objective and Limits of the Review 167 A. Definitions 167 B. Mechanisms of Cooperative Binding 168 C. Methods 170 D. Criteria for Evaluating Kinetic Studies 173 II. Specific Examples 176 A. Hemoglobin 176 B. Glyceraldehyde-3-phosphate Dehydrogenase 184 C. Aspartate Transcarbamylase 193 D. Aspartokinase-Homoserine Dehydrogenase I from Escherichia coli. 199 III. Conclusion 203 References 206

I. Objective and Limits of the Review A. Definitions Allosteric enzymes are defined, for purposes of this review, as those enzymes that can mediate indirect interactions between distinct sites (85). They are often (but not always) characterized by a sigmoidal response of the initial rate to increasing substrate or inhibitor concentration. An increasing number of allosteric enzymes exhibit another type of deviation of the binding curves for ligands from the classical Michaelis hyperbola—namely, a tailing-off at high degrees of saturation (75, 77). Such complex saturation curves can result from the (positive or negative) cooperative interaction between distinct binding sites for the same ligand [homotropic effects (85) ] . Allosteric enzymes seem to be invariably composed of more than one subunit [or protomer (55)]. The oligomeric structure of allosteric enzymes thus provides for the necessary multiplicity of and distinctness of identical sites. Moreover, the modulation of enzymatic activity by metabo­ lites which are structurally unrelated to the substrate bears evidence to the indirect interaction between binding sites for different ligands [heterotropic effects (85) ] . By virtue of these properties, allosteric enzymes are endowed with an unusually versatile potential for regulation of their catalytic activity. Two major questions arise with respect to allosteric enzymes: One bears 167

168

KASPER KIRSCHNER

on the precise regulatory role played by certain key allosteric enzymes when integrated into the complex network of metabolic circuitry. This very important question is intrinsically difficult to answer because the function of allosteric enzymes in the intact cell can hardly be studied directly. This topic will receive only marginal attention in what follows. For pertinent reviews, the recent papers by Sanwal (97), Umbarger (111), Stadtman (102), and Atkinson (5) should be consulted as well as this and the pre­ ceding volumes of Current Topics in Cellular Regulation. The second ques­ tion aims at the molecular mechanisms by which allosteric enzymes interact with substrates and effectors. This problem, which is less complex than the question of function in vivo, is the main topic of this review. The simplified experimental situation with purified enzymes in vitro has the obvious advantage of greater variability and of lending itself to investiga­ tion by direct methods. B. Mechanisms of Cooperative Binding It is useful to distinguish between two aspects of the term "mechanism." Formal mechanisms involve specifying a minimal number of intermediates of defined stoichiometry (but unspecified structure), the interconversions of which suffice to explain the dynamic and equilibrium behavior of chemical reaction systems. In contrast, stereochemical mechanisms provide a detailed three-dimensional description of the structures involved in every elementary reaction step. Of course, the latter is the ultimate goal of understanding chemical reactions in molecular terms. With regard to allosteric enzymes, various formal mechanisms have received detailed theoretical treatment (25, 41, 49, 59, 75-77, 81, 86, 96, 98, 106). It is generally accepted that cooperative binding results from the coupling of binding steps to discrete conformational changes of the protomer. These need not only occur in the isolated protomer but can also be transmitted to other protomers in the quarternary assembly. Thus coupling of the conformation changes can lead to overall cooperativity of ligand binding. The various mechanisms that have been proposed differ mainly in the assumptions about the number of possible states, the degree of coupling [e.g., sequential vs. concerted (71, 76, 116)~], and the relative affinities of the conformational states of the protomer for various ligands (i.e., substrates and effectors). Figure 1 illustrates how a general mecha­ nism involving a tetrameric allosteric protein (33, 75, 76) and only two conformational states can be shown to contain both the sequential (75, 76) and the concerted (85, 96) mechanism as extreme limiting cases. This is not to say that other mechanisms cannot be formulated. Despite the current practice of using squares, circles, and other simple geometrical forms to symbolize discrete conformational states of protomers, the

KINETIC ANALYSIS OF ALLOSTERIC ENZYMES

ffl

169

QP^QQ

if

RI I s Is

ir s Is s| jr si s s s

m

©©

w

1 s1 s 1 s s

FIG. 1. Scheme for the cooperative binding of a ligand to a tetrameric enzyme. An oligomeric protein composed of 4 identical protomers is presented for illustrative pur­ poses. S represents the substrate or any other ligand exhibiting homotropic interactions. Squares and circles indicate different conformational states of the protomer with low and high affinity for the ligand, respectively. Thus, in the absence of ligands, the enzyme will be represented predominantly by the species in the top left-hand corner. Fully saturated enzyme will be mainly in the bottom right-hand form. [From Kirschner (71).] A more general scheme for the tetramer would include a total of 44 species (rather than the 25 species shown). They arise from the binding of ligand either to the squares or to the circles in the 9 hybrid species in the center of the scheme. The mechanism can be expanded readily to oligomers with more than 4 protomers. The species in boldface type represent the two simple limiting cases. The simple sequential mechanism (75, 77) comprises the 5 species on the diagonal with hybrid conformations. The arrows for the interconversion [cf. Eq. (1)] are omitted for the sake of simplicity. The concerted mechanism consists only of the species in boldface type in the left- and right-hand vertical columns (85, 96). The hybrid species are omitted and the two columns are directly connected by reaction arrows (cf. Fig. 2).

mechanisms retain their formal character. This is because hemoglobin is the only example to date of an allosteric protein for which a stereochemical meaning can be assigned to squares and circles (90). It is probably safe to say that the mathematical tools for analyzing the equilibrium behavior [i.e., binding curves and "state functions" (25, 85)"] of quite complex mechanisms are now generally available. The field is in the tantalizing phase where special and general theories are abundant, but decisive experimental data are scarce. In the past, the question of whether or not allosteric enzymes might conform to some unifying mechanistic

170

KASPER KIRSCHNER

principle tended to create two schools of thought (e.g., sequential vs. concerted mechanism). Meanwhile, the discovery of more complex regula­ tory enzymes, such as glutamine synthetase (103), ribonucleotide diphosphate reductase (10), CTP-synthetase (80), and aspartokinase I homoserine dehydrogenase I (17), defies any attempt to find a simple common mechanism. It has also been pointed out (102) that if there were any advantage for regulatory proteins to respond cooperatively to ligands (i.e., substrates and effectors), this property would have been selected for, irrespective of the underlying mechanism. Because it is improbable that any one mechanism will be found to which all allosteric enzymes conform, it is important to learn how large the diversity of mechanisms actually is. It is apparent that more detailed in­ formation is required before new principles of the function and regulation of allosteric enzymes at the molecular level can become discernible. As our understanding of individual allosteric enzymes improves, one can also hope to learn more about general principles of subunit assembly and the forces stabilizing oligomeric complexes. The accumulated experience may prove useful also in exploring the effects of association between dissimilar enzymes (for example, the soluble and membrane-bound multienzyme complexes) on the structure and function of each enzyme involved (46). The purpose of this review is therefore neither to expand previous summaries on the regulatory function of allosteric enzymes in vivo nor to review the mathematical analyses of various hypothetical mechanisms, but rather to survey critically some of the available experimental evidence pertaining directly to the mechanisms of cooperative binding of ligands to certain specific allosteric proteins.

C. Methods Allosteric enzymes are usually identified as belonging to this category on the basis of their complex steady-state kinetics. The substrate saturation (or the inhibition) curves deviate from the normal Michaelis hyperbola. Moreover, most of the information available on allosteric enzymes is in the form of steady-state kinetic data. This is because these studies require only catalytic concentrations of enzyme and because often they can be carried out even in crude extracts. Unfortunately, the information gained from steady-state kinetics is generally insufficient to permit an unequivocal assignment of a detailed formal mechanism. This is particularly true of complex situations as presented by the allosteric enzymes (16). The information is limited to the velocity of rate-limiting steps (Vmax) and to the apparent dissociation constants of substrates (Km) or effectors (Ki) for the enzyme. Only the latter have any direct bearing on the mechanism of

KINETIC ANALYSIS OF ALLOSTERIC ENZYMES

171

allosteric control. In contrast, Michaelis constants are generally complex functions of rate constants (depending on the mechanism) and are identical to true thermodynamic dissociation constants only in limiting cases. There­ fore, the evidence obtained is at best only as good as the information derived from direct binding studies. Moreover, the low concentrations of enzyme (10~10 to 10~6 M) employed preclude the observation of any minor species participating in the overall mechanism. Another serious drawback of the steady-state kinetic approach is that sigmoidal saturation curves are not necessarily reliable evidence for the existence of multiple interacting sites. A number of alternative explanations have been offered, for example, "relaxa­ tion" mechanisms involving single and multiple subunit enzymes (91, 114a>), and the "alternate pathway for two substrate reactions" theory (89,48,104). As allosteric enzymes became available in pure form and in large quan­ tities, the evidence from overall kinetics began to be supplemented by direct binding studies at equilibrium. In favorable cases, the results describe the (usually) noncovalent interactions between substrates (single sub­ strates in multisubstrate reactions or substrate analogs), activators, and inhibitors and the various distinct sites of the allosteric enzyme. The number of binding sites for each ligand often can be directly determined and correlated with the known subunit structure of the enzyme. In terms of the useful nomenclature describing allosteric enzymes as belonging either to the class of "K-systems," "V-systems," or the mixed type (85), direct binding studies focus the attention on the "K-aspect" of allosteric regulation of enzyme activity. Once the thermodynamic relationships have been described in terms of a consistent model, it becomes possible to analyze steady-state kinetic data by assigning specific turnover numbers to the various molecular species present at equilibrium (41 ). Successful as binding studies are in establishing the existence and in estimating the extent of cooperative interactions, the extracted information is again insufficient in principle for an unequivocal distinction between one possible mechanism and another. The validity of a particular model can only be considered proved if the evidence excludes all possible simple mechanisms but one. The limitation of measurements at equilibrium is due to the fact that they record only the overall behavior of complex sys­ tems. Since even the simplest allosteric mechanisms for positive cooperativity (cf. 75, 76, 85, 96) are characterized by at least three parameters, the dilemma arises that a given sigmoidal binding curve can be fitted to a number of different mechanisms with equal success. It has been well documented, particularly in the case of hemoglobin (76, 99), that curvefitting procedures as a diagnostic tool for distinguishing between mechanisms very sensitively depend on the availability of highly accurate experimental

172

KASPER KIRSCHNER

data at the extremes of the saturation curve. The susceptibility of the measurements to various kinds of uncontrolled artifacts recommends great caution in the interpretation of binding data alone. It is also an illusion to believe that various ways of plotting binding data (i.e., degree of saturation versus concentration or the log of concentration of the ligand, reciprocal plots, Hill diagrams, etc.) can do anything to alleviate the dilemma. More­ over, the introduction of additional ligands (giving rise to heterotropic interactions) generally does not assist in defining the mechanism as long as the binding is studied only at equilibrium. What is needed are methods that differ from equilibrium binding studies either by independently measuring parameters related to conformation and/or gross structure of the protein, or by providing another means of measuring independently the individual equilibrium constants involved in the cooperative binding process. Fortunately, a wide spectrum of chemical and physical methods is available today for providing additional information on the allosteric regulation of enzyme activity. X-ray crystallography provides the most direct and detailed evidence on the limiting structures accessible to an enzyme, thus defining the minimal conformational changes occurring during the cooperative binding of the ligand. In the case of hemoglobin it has even been possible to identify with a high degree of probability the protolytic groups responsible for the linkage of oxygen and proton binding (i.e., the Bohr effect) by including structural data from a chemically modified derivative of hemoglobin (90). Highly resolved structures of allosteric enzymes are only just beginning to emerge—for example, aspartate transcarbamylase {117). However, since structure determinations in the crystal can provide only indirect information on the intermediates participating in the cooperative binding of ligands in solution, the results of other methods must be drawn upon to round off the picture. One of these methods is particularly suited for establishing formal mechanisms: fast reaction studies. The high time resolution of the rapid mixing and chemical relaxation techniques allows kinetic measurements to be carried out at substrate concentrations of enzyme (10~~5 to 10~3 M\ i.e., several orders of magnitude higher than commonly employed in steady-state kinetics). Under these conditions, the minor species partici­ pating in the various mechanisms that have been proposed should generally be present in detectable amounts. Furthermore, these concentrations often correspond to the levels of enzymes actually operating in the cell. The principal advantage of kinetics over binding studies at equilibrium arises from the fact that kinetics introduce time as a new coordinate along which the various elementary processes can be separated. In favorable cases, this permits the direct observation of each independent step in a time range characteristic for the particular process. With regard to the more complex

K I N E T I C ANALYSIS O F ALLOSTERIC

ENZYMES

173

allosteric enzymes the number of enzyme-ligand complexes involved and the way in which binding steps are coupled to isomerization processes (conformational changes, for example) can in principle be determined with a minimum of information on the structure of the enzyme. This survey will limit itself to the discussion of a number of allosteric enzymes which have been studied in some detail with the aid of fast reaction methods. It is designed to illustrate the kinetic approach with a few in­ formative examples. The selection is based on the availability of extensive data and on the application of both the stopped-flow and chemical relaxa­ tion techniques. I apologize to those authors whose work has not been mentioned or has not been sufficiently emphasized here. Excellent treatises on the principles, the technological aspects of these methods, and their application to other biological systems are available (13, 34-36, 44, 51, 78). Therefore, only the results obtained with these methods will be discussed. Technological consideration will be more concerned with the spectral properties of the systems under study inasmuch as they permit direct observation of transients or relaxation processes. The restriction of content to kinetic studies is necessary in view of the vast and ever increasing literature on the topic of allosteric enzymes in general. No judgment is passed thereby on the merits of other powerful approaches which focus more on structural features of the interaction of allosteric enzymes and ligands in solution (sedimentation velocity, diffusion, X-ray small-angle and light scattering, nuclear magnetic resonance, electron paramagnetic resonance, optical rotatory dispersion, circular dichroism, and fluorescence, for example). To the contrary, reference to the information obtained by these techniques will be made whenever it can be fruitfully used to comple­ ment the kinetic data or where unresolved controversies await a final resolution. D. Criteria for Evaluating Kinetic Studies Before surveying the results obtained with specific allosteric enzymes, it is helpful to discuss the general conditions under which the performance and evaluation of kinetic studies is meaningful. This is not only appropriate in a paper emphasizing experimental approaches but also serves to caution against the extensive interpretation of insufficient kinetic data or of effects which are prematurely assumed to be intrinsic properties of the enzyme under study. Moreover, these considerations establish the criteria by which the special cases discussed below will be analyzed. 1. PURITY AND STRUCTURE OF THE PROTEIN

Physiochemical studies involving enzymes require that the enzyme preparation be as pure as possible. The concept of purity not only en­ compasses the criterion of homogeneity in polyacrylamide gel electro-

174

KASPER KIRSCHNER

phoresis, but also that of maximal and uniform enzymatic activity. Al­ though the latter quantity is operational by definition, knowledge of it is absolutely essential in the measurement of total concentration of the active (or binding) sites of the enzyme. This is particularly true for allosteric enzymes where desensitization or denaturation can lead to a mixture of physically homogeneous but functionally heterogeneous molecules. For example, binding curves which tail off should be accepted as an intrinsic property of an enzyme (irrespective of the mechanism) only after it has passed stringent tests of physical and functional homogeneity. The concept of purity also encompasses the absence of tightly bound effectors in the stock enzyme solutions. Again the "absence of effector" can be defined only operationally inasmuch as unsuspected ligands can remain undetected. The binding of 2,3-diphosphoglycerate to human hemoglobin (7) is a cautionary tale in this respect. It goes almost without saying that the enzyme should be sufficiently stable under the conditions of the experiment to survive the duration of the measurement. Preferably, the stability should extend over as wide a range of pH, temperature, and ionic strength as possible. For planning kinetic experiments with oligomeric enzymes and for evaluating the data, it is invaluable to know the molecular weight, the number of protomers and their structural identity or nonidentity. If the protein undergoes association-dissociation reactions, the pertinent equi­ librium constants and their linkage to ligand binding must be determined. Furthermore, precise extinction coefficients are required for the reliable determination of total enzyme concentration. Although these prerequisites are necessary for establishing the formal mechanism, there is no upper limit to the additional information required for a detailed description in stereochemical terms. 2. BINDING STUDIES

Accurate equilibrium binding data complement the information on the oligomeric structure of the protein and are indispensable for the planning and interpretation of kinetic experiments. Sigmoidal saturation or inhibition curves as obtained from initial velocity experiments are often taken to reflect binding equilibria, but they can be used for hardly more than a qualitative characterization of the interaction of an allosteric enzyme with substrates and effectors. Equilibrium dialysis is presently the preferred technique for obtaining accurate binding data. The number of moles of ligand bound at saturation per mole of enzyme can usually be extrapolated unequivocally. The necessary controls become more stringent as the dura­ tion of the experiment increases. However, recent advances in technique and/or the manufacture and treatment of dialysis membranes have drastically reduced the time required for equilibration (10, 38, 7^, 93).

K I N E T I C ANALYSIS OF ALLOSTERIC

ENZYMES

175

Binding studies involving changes in fluorescence of the enzyme and/or the ligand or difference spectra are very convenient but must be viewed with caution. This is particularly true with allosteric enzymes where more than one kind of enzyme-ligand complex can exist, each possibly characterized by different spectral coefficients. Moreover, the thermodynamically and kinetically important parameter of the free ligand con­ centration is not readily available from such measurements. It can be calculated only on the basis of assumptions which are difficult to test independently. The dangers of spectrophotometric titration are accentuated when the average thermodynamic dissociation constant is more than one order of magnitude smaller than the concentration of binding sites. Under these conditions, plots of the degree of spectral change versus the total ligand concentration (which constitute the primary experimental data) tend to obliterate any detail at low degrees of saturation. Thus, the existence of positive and/or negative cooperativity can be overlooked, leading to the erroneous conclusion that the binding is hyperbolic. 3. SPECTRAL HANDLES

Since most techniques for the measurement of very rapid reactions are conveniently based on the optical detection of concentration changes, it is practical to study only systems showing finite difference spectra. This restriction seriously limits the number of allosteric enzymes that can be studied by these techniques. However, chromophoric substrate or effector analogs can often be synthesized. As a last resort, recourse can be taken to pH indicators for monitoring reactions via changes of the hydronium ion concentration. While the use of enzymes which have been chemically modified by the introduction of chromophoric "reporter groups" represents another solution to the problem of detecting rapid elementary steps, the specificity of the labeling reaction and the influence of the label on the intrinsic properties of the enzyme must be established by additional control experiments before conclusions about the native enzyme can be made [cf. the recent review by Cohen (19) and references therein]. 4. ANALYSIS OF KINETIC DATA

It is clear that in principle all relevant rate processes must be detected and resolved if a rigorous kinetic treatment of the data is to lead to a unique mechanism. Only then does the kinetic approach develop its full power as a diagnostic tool. While this principle can be circumvented in no way, it is possible that certain processes remain undetected. Escape from detection of processes which should occur can be due to very different reasons for kinetic measurements initiated by stopped-flow or temperature-jump. Trivial reasons are (a) rapidity of reaction, (b) absence of a spectral change or a small enthalpy change for the reaction in question, (c) for-

176

KASPER KIRSCHNER

tuitous overlap of two or more rate processes, and (d) low population of certain intermediates. While reason (a) is less of a problem when the temperature-jump method is used and (b) is serious only if the most rapid process is "silent," (c) and (d) can occur wTith either of the methods. More­ over, the degeneracy can be not only apparent, as in (c) and (d), but also true, i.e., an intrinsic property of the particular mechanism. In evaluating kinetic data, rigorous theoretical treatment of the kinetic consequences of the various possible mechanism is absolutely essential in forming a link between a given mechanism and the data. Closed analytical expressions for various rate processes can be obtained most easily for the kinetic situation as defined by small perturbations of equilibrium [i.e., chemical relaxation (cf. 35, 51, 78)~\, but in practice this can be done only under favorable conditions. In the case of allosteric enzymes, a comparatively slow rate of isomerization with respect to the binding steps constitutes such favorable circumstance. However, in the analysis of most of the results obtained from stopped-flow experiments and for some of the complex chemical relaxation spectra observed after a temperature jump, it will be necessary to use numerical curve-fitting procedures {45, 54-) em­ ploying fast computers. The sacrifice of direct physical reasoning in the analysis of progress curves (i.e., the change of the concentration of a reactant with time) can, to a certain extent, be compensated by the definition of suitable mean relaxation times. This has been done with great success in the related problem of cooperative binding of dye molecules to linear biopolymers (101). The above prerequisites are not all fulfilled in the illustrative examples to be discussed below. However, they serve to define the point of view from which the data are examined in this review and help to focus attention on areas where new information or more rigorous controls and refined measure­ ments are required before a mechanism can be established. II. Specific Examples A. Hemoglobin 1. INTRODUCTION

The binding on oxygen to hemoglobin is the first known example of cooperative binding of a specific ligand to a functional protein. Representing a model for allosteric interactions between different ligands (i.e., oxygen, protons, CO2, and 2,3-diphosphoglycerate) and an enzyme, hemoglobin has been studied more extensively and by a greater variety of methods than any other regulatory protein to date. Fast reaction techniques were applied to the problem of the mechanism of oxygen binding at a very early stage.

KINETIC ANALYSIS OF ALLOSTERIC

ENZYMES

177

In fact, the rapid mixing method was first devised in 1923 specifically for the study of hemoglobin (58). Any attempt to survey comprehensively the vast body of kinetic data which has accumulated since 1923 would exceed the limits of this survey. A recent review by Antonini and Brunori (2) should be consulted for the earlier kinetic studies and for contributions of other methods to the hemo­ globin field. A number of recent papers, in particular the proposal of a stereochemical mechanism by Perutz (90), justify a critical reassessment of the current situation of the kinetics of oxygen binding to hemoglobin. A priori, hemoglobin would seem to be the ideal test case for elucidating the mechanism of cooperative binding. The material is abundantly available and is stable over a wide range of pH, temperature, and ionic strength. The heme group provides for a built in "reporter group" with pronounced difference spectra between oxy- and deoxyhemoglobin. Moreover, a number of iso-electronic ligands (CO, NO, ethylisocyanide) can be used instead of oxygen. The characteristic features of the sigmoidal binding curves (Hill coefficient) and the kinetic progress curves appear to be similar for these ligands. A further advantage is the possibility of separating the a- and ß-chains of hemoglobin. The properties of the isolated chains, when compared to native hemoglobin, provide for a measure of the changes which occur upon formation of the quarternary structure. These must somehow be related to the acquisition of cooperative intersubunit interactions in the tetramer. Many mutant and derivatized hemoglobins are available for studying the effects of single amino acid substitutions and chemical mod­ ifications on its structure and binding behavior. Finally, the atomic models of deoxy and oxy (horse) hemoglobin and of a number of derivatives have been determined at high resolution (90). Notwithstanding the apparently ideal properties of hemoglobin for using fast reaction techniques in elucidating the mechanism of cooperative binding, certain complications make it difficult to interpret the available equilibrium and kinetic data in terms of a self-consistent reaction scheme. First, the component a- and 0-chains are not identical and therefore do not have equivalent domains of intersubunit bonding. Second, there is an equilibrium between αφι tetramere and aß dimers under the usual experi­ mental conditions, the tendency of oxyhemoglobin to dissociate being much greater than that of deoxyhemoglobin (2, 82, 49). However, neither the principal features of the binding curves (i.e., the Hill coefficient n = 2.72.9) nor the kinetic behavior are altered significantly when the conditions are changed from those favoring the tetramer to those stabilizing the oxy dimer (2). This has raised the question whether the functional unit re­ sponsible for cooperative binding is the tetramer or the dimer. Moreover, the discovery of a binding site in deoxy hemoglobin with considerable

178

KASPER KIRSCHNER

affinity for organic phosphates such as 2,3-diphosphoglycerate [and even inorganic phosphate (7)] further complicates the picture. Much of the earlier work has to be reconsidered in the light of this recent finding. In an attempt at simplification, the following discussion of hemoglobin binding kinetics will restrict itself to the results obtained with oxygen as the ligand and where hemoglobin can be considered to be largely in the tetrameric form both in the unliganded and liganded states. It must be em­ phasized that the questions of whether oxyhemoglobin was fully tetrameric under the experimental conditions used and what happens when the conditions are changed so as to favor dimer formation are still contro­ versial (82, 99). For recent studies on the rates of ligand binding at low hemoglobin concentrations, the reader is referred to the recent papers of Gibson and associates (82). A discussion of the interesting effects of organic phosphates on the equilibria and rates of oxygen binding (cf. 88a and references therein) unfortunately exceeds the limits set to this review. 2. RAPID MIXING EXPERIMENTS

When deoxyhemoglobin is mixed with a saturating concentration of carbon monoxide in a stopped-flow apparatus, the rate of CO-hemoglobin formation is autocatalytic (3). This phenomenon may be taken as a kinetic expression of heme interactions. It is interpreted as an approximately 2-fold increase of the association rate constants as the saturation with the ligand proceeds. Since the binding of oxygen to hemoglobin occurs so much faster than that of carbon monoxide it is experimentally very difficult to determine whether the progress curve is also autocatalytic in the former case. This question is still quite controversial (cf. 8, 1+5). For oxygen binding it is possible to fit the kinetic data to the formal Adair scheme, in which the overall process of binding is described as a sequence of four second-order steps (43, 95) : Hb4(02)„_i + 0 2 ^ Hb4(02)n

with n = 1 to 4

(l)

kn

This mechanism is equivalent to the simple sequential mechanism (cf. Fig. 1) (75, 77), which also comprises 5 different molecular species inter­ related by four overall binding steps. Recently, the kinetics have been reinvestigated by Gibson (45), using refined methods of data acquisition and analysis. Since the progress curve does not permit the direct observation of the individual elementary steps involved, additional information was needed to restrict the number of possible solutions for the set of 8 rate constants. The auxiliary data were obtained from independent measure­ ments of the association and dissociation rate constants of the last step (W and &4, respectively), from concentration-jump relaxation experi-

179

KINETIC ANALYSIS OF ALLOSTERIC ENZYMES

ments and from binding isotherms. A simultaneous fit of all these data to the Adair scheme (Eq. 1) has led to a unique set of 8 rate constants (Table I). It can be seen from Table I that the association rate constants (which determine the overall rate of complex formation in the presence of high oxygen concentrations) actually increase 2-fold as the reaction proceeds. However, the progressive changes are not monotonie. The increase of affinity, which is the basis for the overall cooperative binding curve, is attributable mainly to a marked decrease of the dissociation rate constants as the degree of saturation increases. These values also agree with the early finding that the overall kinetics of dissociation of oxyhemoglobin (in the presence of dithionite as a scavenger for oxygen) is characterized by an initial lag before becoming first order (25ay 58, 96a). The earlier interpretation that this is due to a limitation of the overall rate by the first oxygen coming off appears to be correct to a first approximation. It has been pointed out by Gibson (45) that the quantitative fit of his kinetic data to the Adair scheme does not prove this mechanism to be correct. This is mainly because the rather monotonie progress curve does not yield any independent information on the total number of elementary steps (or intermediates) involved in going from deoxyhemoglobin to oxy­ hemoglobin. It is noteworthy that a correlation of data from equilibrium binding curves and stopped-flow kinetics is possible only if the predominant intermediates are the same in both experimental situations. If the structural changes which must form the molecular basis of cooperative binding were slower than the binding steps, this assumption would no longer be valid. An example for this situation is yeast glyceraldehyde-3-phosphate dehydrogenase (see discussion of this enzyme in Section II, B) (74). This is prob­ ably not true for hemoglobin since stopped-flow experiments at high oxygen TABLE I RATES AND EQUILIBRIUM CONSTANTS FOR THE BINDING OF OXYGEN

TO

HUMAN

HEMOGLOBIN A

PHOSPHATE B U F F E R pH

a 6

fcn

IN

0.1

M

7.0 AND 21.5°. e · 6 iCn

n

(μΜ _1 sec-1)

(sec-1)

1 2 3 4

17.7 33.2 4.9 33.0

1900 158 539 50

KTi

Kn/iCn

(μΜ) 107.0 4.8 110.0 1.5

From Gibson {^5). The symbols for the rate constants correspond to Eq. (1).

ISO

KASPER

KIRSCHNER

concentrations (8) have shown that no first-order process (i.e., a structural change) becomes rate limiting at reaction half-lives down to 1 msec. If the Adair or simple sequential mechanism were correct, the structural changes would be triggered by the binding of oxygen and would occur concomitantly with the binding. Therefore, they might not have to be included as inde­ pendent elementary steps. The values of the rate constants obtained by Gibson {45) are not easily reconciled with the predictions of the concerted mechanism. The alternating changes of the rate and equilibrium constants (cf. Table I) with increasing degree of saturation appear to be in agree­ ment with the general idea {2) that the aß dimers contribute much of the overall cooperativity of ligand binding by hemoglobin. Apparently the progress curves show no evidence of a dissociation of oxyhemoglobin tetramere to the oxy dimers. Although deoxyhemoglobin is probably completely tetrameric under Gibson's {45) experimental condi­ tions {82, 69), this process must occur to a certain extent in oxyhemoglobin after it has been formed from deoxyhemoglobin. It remains an open question whether the effects wrere so small (or so rapid) that they could not be ob­ served. It is also still controversial, whether the dissociation of oxyhemo­ globin to dimers and monomers is accompanied by spectral changes {84) or not {89). 3. CHEMICAL RELAXATION EXPERIMENTS

In view of the high rates of oxygen binding to hemoglobin and of the isomerization steps involved, the different approach used in the tempera­ ture-jump method appears to be well suited a priori for a resolution of the mechanism into the underlying elementary steps. Moreover, the high time resolution of this technique permits kinetic studies at high hemoglobin concentrations where the dissociation of oxyhemoglobin to dimers can be minimized. Such experiments have been carried out recently with sheep hemoglobin in dilute borate buffer pH 9.1 {11, 62, 99). Similar experiments have been carried out with human hemoglobin at neutral pH [unpublished work of M. Brunori cited in {2) and T. M. Schuster, personal communica­ tion] and have led to similar relaxation spectra. As was to be expected, the relaxation spectrum is more complex than the progress curve obtained from stopped-flow kinetics {45). At least 4 welldefined relaxation processes can be separated. The fastest process could only be resolved with the aid of the electrical field pulse method {62). The relaxation time (τ 0 ^70 nsec) characterizes a first-order process which apparently is not linked directly to the conformational changes accom­ panying ligand binding. The peculiar dependence of the amplitude of the fast process on pH suggests (but does not prove) that it might be associated with the Bohr effect (i.e., the linkage of proton and oxygen

K I N E T I C ANALYSIS O F A L L O S T E R I C

ENZYMES

181

binding). The somewhat slower processes characterized by n (1-5 msec) and r2 (5-50 msec) are clearly linked to oxygen binding steps. The slowest process observed (r3 = 5-50 msec, depending on protein concentration, see below) can be interpreted in terms of the tetramer-dimer equilibrium of hemoglobin which is linked to oxygen binding {49). However, since the amplitude of the slow relaxation process vanishes under conditions favoring the tetrameric state of oxyhemoglobin (low ionic strength and high protein concentration), τχ and r 2 appear to be the only relaxation times pertaining to the mechanism of cooperative binding of oxygen to tetrameric hemo­ globin. This requires the existence of at least one intermediate at detectable concentrations between the unliganded and liganded forms of hemoglobin. Moreover, the relaxation spectrum is similarly simplified to only n and τ2 when the conditions are changed so as to promote the dissociation of oxy­ hemoglobin to dimers (i.e., in strong salt and at low protein concentration). This finding represents strong evidence in favor of the "functional dimer hypothesis" (see 2). Schuster and Ilgenfritz (99, 100) have analyzed the data obtained with tetrameric hemoglobin by comparison with theoretical calculations based on the four-step Adair scheme and the concerted mechanism (cf. Figs. 1 and 2). Unfortunately, the failure to separate all the relaxation processes to be anticipated for any of the proposed mechanisms does not allow the derivation of closed analytical expressions relating the reciprocal relaxation times to rate constants and equilibrium concentrations of the reacting species. Using accurate data on the binding isotherms of sheep hemoglobin at various temperatures and Gibson's (43) earlier set of rate constants as starting values, the concentration dependences of the reciprocal relaxation times and the corresponding amplitudes of all possible relaxation processes were calculated. The Adair or simple sequential scheme is described by four relaxation times. For the case of the concerted mechanism the assump­ tion was made that the concerted structural transitions should be faster than the binding steps and that they do not give rise to spectral changes per se. As discussed above, the first assumption appears to be justified, whereas the second assumption is controversial (84, 89). As a consequence, the total of nine relaxation processes anticipated for the concerted mecha­ nism (33, 34) is divided into a class of 5 rapid (and undetectable) isomerization steps and another class of comparatively slow relaxation processes involving 4 binding steps coupled to each other via the rapid isomerization reactions. The interesting result is that the relaxation spectra calculated for both mechanisms are very similar. The calculations predict that only two (at the most three) out of a maximum of four relaxation processes would be observable in each case with the available instrumental sensi­ tivity. This is a direct consequence of the high degree of cooperativity of

182

KASPER KIRSCHNER

oxygen binding to hemoglobin, leading to low concentrations of the inter­ mediates throughout the saturation curve for each mechanism. The situa­ tion is analogous to the equally good fit obtained to the same saturation curve with both mechanisms, for the same reasons. Although the chemical relaxation data qualitatively agree with the predictions of both mechanisms (and therefore cannot distinguish between them), the relative proportion of the amplitude of the rapid process characterized by n is compatible with neither one. In conjunction with Gibson's {46) analysis, it can be said that the mechanism probably requires some expansion of the sequential mechanism. While the concerted mechanism {86, 96) seems to be clearly ruled out, it is possible that the true mechanism may contain elements of sequential and concerted structural changes. Exactly this type of mechanism has been discussed by Perutz {90). 4. DISCUSSION

It is necessary to correlate the kinetic data with the wealth of additional information available on various aspects of structure-function relationships in hemoglobin. As Antonini and Brunori {2) point out, it is not possible to explain all known properties of hemoglobin by a unique mechanism. How­ ever, two fundamental controversies appear to be nearing their reconcilia­ tion in the light of more recent information. One is the question of whether the functional unit of hemoglobin is the aß dimer or the a2ß2 tetramer. The other question concerns the manner of coupling of the tertiary and qua­ ternary changes (sequential or concerted) which form the molecular basis of cooperative binding of oxygen to hemoglobin. Both aspects are obviously interrelated but will be discussed separately for the sake of convenience. The kinetics observed after rapid mixing of oxygen and hemoglobin cannot be fitted to a dimer model {46). This is also true for the binding curve, since the Hill coefficient is close to the value of 3. Recent ultracentrifugal studies {32, 69) show that deoxyhemoglobin is a stable tetramer even at high dilution (if4,2 ^p M.W. 4 0 0 , 0 0 0

monomers M.W. 100,000

FIG. 1. Molecular forms of muscle phosphorylase.

essentially inactive in the absence of AMP, and phosphorylase a, a tetramer of M.W. 400,000, active in the absence of any effector provided it is fully saturated with substrate (Fig. 1). Conversion of phosphorylase 6 to a occurs through phosphorylation of the protein by Mg-ATP in a reaction catalyzed by phosphorylase kinase (89) ; phosphorylase a is converted back to b by phosphorylase phosphatase (26,54,84,85). Both phosphorylases b and a can be reversibly dissociated into monomeric subunits on treatment with sulfhydryl reagents such as p-CMB* (106) or DTNB (30, 82) or "deforming agents" (14-7) such as imidazole citrate. Some of the physical constants of the enzyme are summarized in Table I. From sedimentation equilibrium studies carried out in a number of de­ naturing solvents or polyacrylamide gel electrophoreses in sodium dodecyl sulfate, no evidence could be obtained that the 100,000 M.W. subunit (17, 14S) is made up of more than a single polypeptide chain (144) ; furthermore, no chemical data supports the view that the two subunits of phosphorylase b might be of more than one type as recently suggested by electron microscopy (159). As commonly found for regulatory enzymes, phosphorylase contains a multiplicity of sites, all affecting the activity of the enzyme (Fig. 2). First, there is a catalytic site(s), binding the three substrates (glycogen, Pi and glucose-1-P) and some inhibitors (e.g., glucose, gluconolactone, or * Abbreviations: p-CMB, p-chloromercuribenzoate; DTNB, 5,5'-dithiobis(2-nitrobenzoic acid) ; PLP, pyridoxal 5'-phosphate; UDPG, uridinediphosphoglucose; 5'-P-Pxy, 5'-phosphopyridoxyl; FDNB, fluorodinitrobenzene; EDTA, ethylenediaminetetraacetic acid; EGTA, ethyleneglycol bis (/3-aminoethyl ether)-iV^iV'-tetraacetic acid; KAF, phosphorylase kinase activating factor.

Α*οι%

β

Xh

0.53

13.1 dzO.2

110 X 65 X 55 109 X 62 X 55

—

—■

—

—

Refractometry and amino acid analysis

Electron microscopy Small angle x-ray scattering

Gel electrophoresis in SDS Sedimentation equilibrium in 6 M guanidinium«Cl (v = 0.737)

Measurements with the native proteins were carried out in 50 TCIM glycerophosphate, pH 7.0.

9 260:280 absorbance ratio

8

7 Molecular dimensions (A) IXW

(17)

(17)

(17, 11, 52)

(14) (129a)

(17,144) (156)

(17, US)

(34)

Pycometry and Linderstr0m-Lang columns

(34)

Gel filtration Calculated from properties 1 and 5 Gel filtration

(17)

Reference

Analytical ultracentrifugation

Technique

400,000 d= 2 % Sedimentation equilibrium

100,000 97,000

200,000 dz 1%

5 Molecular weight

6 Subunit molecular weight

0.746 dz 0.002

6 3 . 0 db 1.7

3.39 3.36

4.33 4.29 49.3 ± 0 . 9

13.7 z b 0 . 0 5

8 . 8 dz 0 . 0 5

Phosphorylase ò Phosphorylase a;

4 Apparent specific volume

3 Stoke's radius (A)

2 D20,w

1 «20, w

Property

TABLE I

PHYSICAL PARAMETERS OF R A B B I T MUSCLE PHOSPHORYLASE 0

to1

w

Ö

P

H

P

X

**3

8

tì

H-

CONTROL OF GLYCOGEN DEGRADATION

215

FIG. 2. Schematic representation of sites on phosphorylase a monomer.

U D P G ) . Second, there is a nucleotide binding site which is the site of allosteric control: binding of AMP produces the change in conformation (157) resulting in the appearance of enzymatic activity in phosphorylase b. The same site binds ATP (105, 123) which acts as an allosteric inhibitor, whereas glucose-6-P (also an inhibitor under certain conditions) most probably binds at a separate site. Third, there is the site phosphorylated during the 6 to a conversion; phosphorylation freezes the molecule in the active conformation and provides for a covalent control of enzymatic activity in addition to the allosteric control mentioned above. Fourth, there is the site binding pyridoxal 5'-phosphate (PLP) which is indispensable to the activity of the enzyme. Finally, there must be several secondary sites or distribution of groups involved in the subunit assembly of the molecule and responsible for the formation of the dimeric and tetrameric forms of the enzyme. While nothing is known about the general distribution of these sites on the surface of the enzyme itself, a number of definite homotropic or heterotropic interactions have been recognized to the extent that inter­ action at any one site appears to affect the properties of all the other sites and therefore, the general behavior of the molecule. These will be described in more detail later. Relatively little is known on the chemical properties of phosphorylase; clearly, a detailed understanding of its structural features will have to await its complete sequence analysis and three-dimensional structure by X-ray crystallography. No amino-terminal group, including acetylated or formylated residues, was detected in any phosphorylase so far examined

216

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

(#).* More unusually, no carboxyl-terminal group was found in the rabbit muscle enzyme; on the other hand, a COOH-terminal lysyl-isoleucine sequence was found in rat phosphorylases {145). Phosphorylase possesses nine free SH groups (no SS bond) {5), some of which are intimately linked to the state of aggregation, or the activity of the enzyme: (a) substitution of two fast reacting SH groups per monomer by reagents such as DTNB {82), FDNB {52), iodoacetamide {4, 6), etc., has little effect on the catalytic or oligomeric properties of the enzyme except that homotropic interactions are abolished {82). In other words, one observes a conversion from allosteric to Michaelian kinetics, (b) A slower reaction with 1-2 additional SH groups results in dissociation of the protein to the monomeric state with complete abolition of enzymatic activity. (c) Complete substitution of all nine sulfhydryl groups is attained only after digestion with pepsin {5). Ligand or covalent-induced conformational changes dramatically alter the pattern of sulfhydryl reactivity: for instance, phosphorylase a lacks the two slow-reacting sulfhydryl groups seen in phosphorylase b {136). Likewise, AMP converts the reactivity of the SH groups of phosphorylase ò to that of phosphorylase a {30). B. The Pyridoxal 5'-Phosphate Site and Role of the Cofactor Pyridoxal o'-P (PLP) has been found in stoichiometric amounts (one molecule per enzyme subunit) in all phosphorylases so far isolated. Par­ ticular interest in the role of this cofactor was stimulated by the finding that whereas its removal from phosphorylase led to total inactivation of the enzyme {80), reduction of the protein with sodium borohydride, which fixes the cofactor to the protein through its formyl group, results in a material which is still enzymatically active {40, 43). All other classical B 6 enzymes in which pyridoxal phosphate is directly involved in catalysis are totally inactivated by this kind of treatment {39, 57). One is, therefore, left with the following alternatives: (a) PLP partici­ pates directly in catalysis, but then a functional group other than the 4-aldehyde must be involved, (b) Phosphorylase is a double-headed enzyme capable of catalyzing a second enzymatic reaction (e.g., one of the classical reactions of PLP-containing enzymes), but none was detected {61). (c) The cofactor is not directly implicated in catalysis, but can be used to control the activity of the enzyme. Against this hypothesis are two arguments: (i) Control mechanisms and effectors of enzyme activity usually vary considerably from species to species and even tissue to tissue. Yet, PLP has been found in phosphorylases from mammals, avians, * Recently, the NH2-terminal sequence of dogfish muscle phosphorylase was shown to be: N-acyl-Ser-Lys-Pro-Lys-Ser-Asp-Met (P. Cohen, unpublished results).

217

CONTROL OF GLYCOGEN DEGRADATION

crustaceans, bacteria, and higher plants, (ii) In order to control the activity of phosphorylase through the cofactor, one should be able to either remove it, modify it, or otherwise alter the site at which it is bound, but PLP is very strongly bound to the enzyme and essentially buried within the protein molecule, (d) PLP is involved neither in direct catalysis nor control but is present as a highly specific structural determinant essential for enzymatic activity. However, in terms of cellular metabolism and no matter how crucial this role may be, it would still be an expensive way to maintain a protein in the proper conformation, and one would wonder why certain species or organisms, after countless mutations, would have not been able to find a more conventional way of doing so. To make things worse, in rabbit muscle, for instance, there appears to be more vitamin B 6 stored in glycogen phosphorylase than in all the other classical B 6 enzymes taken together (91 ) . Originally, on the basis of unusual spectral characteristics of the native enzyme (absorption band at 330 nm) and its resistance to sodium borohydride reduction, a substituted aldamine structure wTas postulated be­ tween the cofactor and the protein (86). Recent evidence indicates, how­ ever, that PLP is indeed linked as a Schiff base but that the unusual properties mentioned above result from the fact that the cofactor sits in a hydrophobic pocket (27, 28, 80a). Limited tryptic and chymotryptic hy­ drolysis together with CNBr cleavage of NaBH 4 -reduced phosphorylase led to the following sequence for the PLP-binding site (Fig. 3) (4-7). Several features emerge from this structure: (a) the PLP binding site is almost free from basic amino acids; none occur over a sequence of about 35 residues, while statistically the enzyme contains one basic amino acid every 7-8 residues (146). By contrast, the phosphorylated site of the 1

5

10

15

. . . Arg-VaI-Ser-Leu-AIa-GIx-Lys-VaI -11e-Pro-AIa-AIa-Asp-Leu-Ser-

20

25

30

GI x-G Ix-1 Ie-Ser-Thr-AIa-GIy-Thr-GIn-AIa-Ser-GIy-Thr-GIy-Asp-

35

Met-Lys(5'-P-Pxy)-Phe-Met-Gly-Arg-Thr-Leu(Glx,Asx,Thr)

-Met

FIG. 3. Amino acid sequence of the pyridoxal o'-phosphate binding site in rabbit muscle phosphorylase. *In dogfish muscle phosphorylase, the following COOH-terminal sequence for the PLP-binding site was found: Met-Gly-Arg-Thr-Leu-Gln-Asn-Thr-Met.

218

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

enzyme is extremely basic (see below). The PLP-binding site in Escherichia coli arginine decarboxylase also appears to be remarkably poor in basic amino acids (7). (b) There is an arginyl side chain 4 residues removed from the P-pyridoxyl-lysyl group that could perhaps be involved in binding the 5 r -phosphate of PLP and provide the correct positioning of the cofactor required for activity (47). E. coli arginine decarboxylase has a histidyl group also four residues away that might serve the same function (7). (c) As noted with E. coli glutamic acid decarboxylase, there is an alterna­ tion of hydrophilic and hydrophobic residues (at least ten) preceding the P-pyridoxyllysyl group suggesting that perhaps this segment of the mole­ cule exists in a /^-conformation {151). In fact, the proximal (aminoterminal) portion of the PLP-binding site is remarkably hydrophilic while the amino acids distal to the cofactor are always hydrophobic, an observation also made in E. coli glutamic acid {151), lysine {137), and arginine decarboxylases (7). (d) The Phe-Met bond distal to the P-pyridoxyllysyl residue appears to be split both by trypsin and chymotrypsin; a similar observation was made with E. coli glutamic acid de­ carboxylase which possesses an identical P-pyridoxyllysyl-phenylalanine sequence {151). The prosthetic group in phosphorylase is "buried" within the protein and thus inaccessible to ordinary carbonyl reagents. Its removal requires a prior distortion (deformation) of the molecule that exposes the cofactor {63, 147) ; when this is carried out under mild conditions (e.g., imidazolecitrate buffer at pH 6.0, 0°) in the presence of certain carbonyl reagents (e.g., L-cysteine), resolution is completely reversible. Deformation of the enzyme is accompanied by dissociation to monomers, with loss of enzymatic activity; with resolution, the fluorescence emission at 530 nm due to the bound cofactor is lost {63). Resolution seems highly specific toward both the nature of the deforming agent and that of the carbonyl reagent: it proceeds readily with L-cysteine, but not with homocysteine, cysteamine, dimercaptopropanol, etc. Most surprising, it does not proceed with D-cysteine, an indication that the reaction is stereospecific. It is also prevented by phosphorylation of the protein (conversion of phosphorylase ò to a) or addition of AMP, empha­ sizing once more the vast changes in conformation that occur under these conditions {148). The apoenzyme is totally inactive, but readdition of PLP restores full activity; reconstitution is strongly temperature-dependent with an energy of activation of ca. 22 Kcal/mole demonstrating the con­ siderable changes in conformation that occur {62,148,149). The apoenzyme has a high affinity for the cofactor and will preferentially remove this compound when it is mixed with many of its analogs. Both pyridoxal and 5-deoxypyridoxal or the pyridoxal 5'-phosphate monomethyl ester {65)

CONTROL OF GLYCOGEN DEGRADATION

219

(and, for that matter, other aldehydes unrelated to PLP) also form Schiff bases with the enzyme and restore both the state of association and allosteric properties of the native protein, but not its catalytic activity (149)· Since NaBH 4 -reduced phosphorylase is enzymatically active, a detailed study was undertaken to assess the possible participation of other func­ tional groups of PLP in catalysis. To this effect, analogs modified in every single position around the pyridine ring were tested for their ability to reactivate apophosphorylase (127,149). Table II summarizes some of these results, indicating that positions 2, 3, and 6 of PLP are not essential for catalysis. The potential aldehyde group in position 4 is necessary for the binding of the cofactor, but not for enzymatic activity. Most analogs of PLP modified in position 5, such as pyridoxal 5'-sulfate, 5'-acetic acid, or the 5'-phosphate monomethyl ester are inactive, but not the 5'-methylenephosphonate derivative (74) which reactivates the enzyme up to about 20% (160), indicating that a group with a pK around neutrality might be required for activity. This possibility seemed further supported by the finding that with decreasing pH, there is a proportional loss of activity and increase in fluorescence quantum yield of the bound PLP, with both processes showing an identical pK of 6.2 (27, 28). However, the phosphate group of PLP cannot be directly implicated, since pyridoxal 5'-P monomethylester (which lacks a group with pK approximately 6.2) displays a similar fluorescence titration curve; this latter effect must therefore be ascribed to an alteration of the structure of the enzyme (88). There is, of course, no exchange of the 5'-phosphate group of PLP with either Pi or glucose-1-P, two of the substrates of phosphorylase (80). On the basis of differential spectroscopic data, it was recently proposed that the pyridinium nitrogen of P L P could be implicated in catalysis by forming an ion pair with the negatively charged groups of the substrates, Pi or glucose-1-P (8). While this hypothesis could not be substantiated, no analog of PLP modified in the 1 position has been shown unambiguously to activate apophosphorylase; therefore, the possibility still remains that activity depends upon the integrity of the pyridine ring nitrogen; a proton transfer between this atom and the 5'-phosphate or phosphonate group could be envisaged. C. The Phosphorylated Site Phosphorylation of a single seryl residue by Mg-ATP and phosphorylase kinase brings about the conversion of phosphorylase & to a in which the enzyme is covalently fixed in the active conformation (89). All phosphorylases so far investigated appear to have very similar sequences at their phosphorylated site (Table III) ; these bear no resemblance to sequences

—

—

9.0

8

Me

6

8.2 8.2 8.2

PL PL 5'-sulfate a6-Pyridoxal acetic acid

9.2

25

Méthylène phosphonate

5

6.5 6.9 7.7

Pyridoxine 5'-P Pyridoxamine 5'-P Pyridoxic acid 5'-P

—

8.1

60

Reduced phosphorylase 6

4

8.5

7.5

ω-Me-PLP

(«20,w)

Aggregation state

ΛΓ-Me-PLP

Modification

Nonactive analogs

—

25

0 Me PLP

3

8.3

8.4

(S20,w)

Aggregation state

(PLP)

—

65

100

Nor-PLP

Control

Modification

% Specific activity

Active analogs

WITH ANALOGS OF PYRIDOXAL 5'-PHOSPHATE

2

1

Position modified

TABLE II

CHEMICAL AND PHYSICAL PROPERTIES OF PHOSPHORYLASE b RECONSTITUTED

s w

Ö

>

S3

*< a

F

m

GO O

*4

w

to

LO

Argr-Gln-Ile-Ser (P)-JZe-Arg

Glu-Ar0-Arg-Lys-Gln-Ile-Ser (P) -Val-Arg-Gly-Leu

Ser-Asp-Gln-Asp-Lys-Arg-Lys-Gln-Ile-Ser (P) -Val-Arg-Gly-Leu

(168)

b

(146)

(73)

(121)

Ser-Asp-Gln-Glu-Lys-Arg-Lys-Gln-Ile-Ser (P) -Val-Arg-Gly-Leu Lys-Gln-Ile-Ser (P) -Val-Arg

References

Amino acid residues

Amino acid residues that differ from those of the rabbit muscle phosphopeptide are italicized. ' P. Cohen, unpublished results.

1

Rabbit liver

Dogfish skeletal muscle

Rat skeletal muscle

Human skeletal muscle

Rabbit skeletal muscle

Tissue

STRUCTURE OF THE PHOSPHORYLATED SITES OF VARIOUS PHOSPHORYLASES 0

TABLE III

to to 1-^

O 525

Ö î> H

>

G

Ö

G O G H

O *4 G

50 O F

o

G

222

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

obtained from the active site peptides of "seryl" proteases or esterases or from other enzymes that can also be phosphorylated (alkaline phosphatase or phosphoglucomutase). The phosphoseryl residue is flanked by two hydrophobic amino acids followed by an arginyl residue on the distal side; it must belong to a highly basic region of the enzyme since CNBr fragmenta­ tion of 32P-labeled phosphorylase a leads to the isolation of a large radio­ active peptide (approximately 88 amino acids) with an isoelectric point of 10.5 and which contains on the average one basic residue for every 5 or 6 amino acids (186). The phosphorylated site must occupy an exposed position on the surface of the protein and contain some structural features necessary for a double recognition by both phosphorylase kinase and phosphatase. It is also readily attacked by several proteolytic enzymes; limited proteolysis by trypsin removes a phosphorylated hexapeptide ( Lys-Gln-Ile-SerOP-Val-Arg) (41, 121) and the remainder of the molecule, which dissociates to a dimeric species designated as phosphorylase b', is still enzymatically active provided AMP is added. Of course, it can no longer be reconverted to phosphorylase a. As expected from the similarities in the structures of the phosphorylated sites among various phosphorylases, both phosphorylase kinase and phosphatase show a low degree of organ or species specificity in that enzymes from rabbit skeletal muscle will act on rabbit heart or liver phosphorylases or on phosphorylases from human (171), rat (146), frog (111), lobster (29), and dogfish muscle (17) [but not on yeast phosphoryl­ ase which also undergoes a ò to a conversion (46) ] . On the other hand, as expected from enzymes having an important regulatory function, they display a high degree of substrate specificity restricted to the phosphorylase molecule, as will be discussed in Sections IV, A and V. D. Enzymatic Properties In spite of extensive studies on the mode of action of phosphorylase, no detailed information exists as to the exact mechanism of catalysis and the nature of the residues involved. Glycogen phosphorylase catalyzes the reversible reaction: a-D-Glucose-l-P + glycogen (n ) ^ Pi + glycogen (n+1)

where n represents the number of glucosyl residues in the polysaccharide. Equilibrium is reached at a Pi/glucose-1-P ratio of 3.6 at pH 6.8, indicating that glycogen synthesis is slightly favored; since Pi is a stronger acid than glucose-1-P, the equilibrium is pH dependent (28, 26). Yet, in spite of this, several observations support the view that under physiological conditions, this enzyme functions almost entirely in the direction of glycogen break­ down: (a) physiological conditions leading to an increase in phosphorylase

CONTROL OF GLYCOGEN DEGRADATION

223

activity, such as the administration of epinephrine or glucagon, always brings about a breakdown of glycogen (81); and (b) patients suffering from McArdle's disease, a hereditary myopathy in which phosphorylase is missing (117,140), or I-strain mice (102) deficient in phosphorylase kinase, generally show an accumulation of glycogen. A separate pathway for glyco­ gen synthesis was suggested by the finding that this reaction occurs even at a high Pi/glucose-1-P ratio that should favor glycogen breakdown (97). In keeping with other phosphorolytic reactions, it is the C ( l ) - 0 bond that is cleaved in both glucose-1-P and the a (1—>4) glucosidic linkage (18). The reaction proceeds with absolute retention of configuration, and only in the presence of oligosaccharide primers (1) of at least 3-4 glucose units. No exchange was detected between Pi and glucose-1-P in the absence of glycogen, and no exchange between free glucose and either glucose-1-P or glycogen was found (19). Similar studies provided unequivocal evidence for a double displacement mechanism in sucrose phosphorylase (53, 162), and the glucosyl enzyme intermediate was ultimately isolated. Recently, a specific inhibition of phosphorylase by 1,5-gluconolactone (competitive toward glucose- 1-P and noncompetitive with respect to glycogen when assays were made in the direction of glycogen synthesis) has been described; it was attributed to interaction of the inhibitor with the glucosyl transfer site of the enzyme and interpreted in terms of a transition state involving the formation of an enzyme glucosyl complex in which the glucosyl residue would occupy (like 1,5-gluconolactone) a half-chair conformation (155). That the transition state might also involve the formation of a carbonium ion was further suggested by the finding of secondary isotope effects in reaction mixtures containing 1-deuterio-a-Dglucose-1-P. Monod first predicted that phosphorylase would behave like other regulatory (allosteric) enzymes and display cooperative kinetics (118). The allosteric constant L, describing the equilibrium between the active and inactive states of phosphorylase, has been determined: at 23° (measured in the absence of other ligands) it is 2100 for phosphorylase b and 3-13 for phosphorylase a (83). This is in keeping with the observation that AMP is indispensable for the activity of phosphorylase ö but merely stimulates phosphorylase a. This nucleotide was shown to bind to phosphorylase ò in a cooperative fashion with a Hill coefficient of nearly 2 (105). Since the active conformation of phosphorylase b is analogous to that obtained by phosphorylation of the protein, both can be looked upon as parallel cooperative systems, differing only in the details of their allosteric behavior. The situation, of course, is not that simple since effectors such as glucose-6-P and ATP which are potent inhibitors of phosphorylase 6 have no effect on the activity of phosphorylase a. Both active forms have a strong tendency

224

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

to tetramerize to the extent that, originally, activity was thought to depend on this change in quaternary structure. This view cannot be main­ tained today. First, rabbit muscle phosphorylase a dissociates from tetramer to dimer in the presence of glycogen, glucose or high salt concentrations with concomitant increase in specific activity {112, 164). Similarly, the tetramerization of phosphorylase b produced by AMP does not occur under assay conditions (high dilution) and, therefore, is not a prerequisite for activation {157). Second, there is no consistent relationship between phosphorylation and tetramerization: whereas phosphorylases from rabbit, human, rat, and frog muscle (as well as that from human platelet) tetram­ erize readily, others including phosphorylases from rabbit heart (isozyme 1), from dogfish and lobster tail muscle and from rabbit liver do not. As for yeast phosphorylase, the contrary situation prevails in that the phosphorylated form of the enzyme exists predominantly as a dimer while the inactive b form is tetrameric {48). Actually, one does not really know whether any of these changes in quaternary structure are of physiological significance since conditions under which the enzyme exists intracellularly [it appears to be bound to glycogen particles {113)~\ differ vastly from those that prevail in vitro {60). There is no evidence that the monomeric form of rabbit muscle phosphorylase a or ò possesses any intrinsic enzymatic activity. On the other hand, phosphorylases from the insect flight muscle {15) and Tetrahymena pyriformis {80b) have been reported to exist in the monomeric state. Many compounds have been shown to affect the activity of phospho­ rylase; those of possible metabolic significance will be discussed in Section III. They are all capable of homotropic and heterotropic interactions; in fact nearly all the binding sites interact with one another to produce heterotropic effects that have been summarized in Table IV. For instance, the affinity of phosphorylase b for AMP is increased by two orders of magnitude (from a KdiSS = 5 X 10~3 to 8 X IO -5 M) by increasing con­ centrations of glucose-1-P and Pij conversely AMP increases considerably the affinity of the enzyme for substrates, and it is largely on this basis that the AMP activation of phosphorylase 6 is explained {68, 83, 100). Phos­ phorylase, like many other regulatory proteins, can be desensitized by treatment with various agents, e.g., limited tryptic attack {55) or reaction with sulfhydryl reagents {82) or glutaraldehyde {165). The desensitized enzyme no longer displays homotropic cooperativity but still requires AMP for activity. UDPG, which acts as a competitive inhibitor toward the substrate, actually leads to an increase in activity of the enzyme when present at low concentrations {105, 108). Such an effect was initially described for

Required for 6 activity. Promotes active conformation. Heterotropic co­ operativity with Pi, G-I-P

Freezes enzyme in active conformation, abolishes substrate cooperativity

Indispensable for activity

Monomers are inactive; dimer a more active than tetramer a

AMP

Seryl phosphate

PLP

Aggregation Binds to all forms of aggre­ gation

Not required for binding of AMP

Increases binding by 25-fold; abolishes cooperativity.

Homotropic inter­ actions on b, none on a

Heterotropic inter­ action with Pj and G-I-P

AMP PLP

Inhibits resolution in the absence of glycogen

Inhibits resolution, not reconstitution

Slight inhibition of resolution by G-I-P, Pi, U D P G

Resolution greatly No major effect on enhanced by phosphorylation or monomerization dephosphorylation

Not required for None phosphorylation or dephosphorylation

None

Inhibits dephosphorylation

Glycogen stimulates phosphorylation (may act on kinase rather than phosphorylase)

Seryl phosphate

Additional aggre­ gation site(s) become functional when primary ones are blocked

Maintains quaternary structure

Induces active conformation and aggregation to tetramers

Induces active conformation and aggregation to tetramers

Glycogen promotes dissociation of tetramer a to dimer b

Aggregation

3

o

H

a o w > >

O O 2 U

tr o

o o1

o

o

o

This table is meant to be read in one direction only: it summarizes the effects of the sites printed in boldface (left) on those at the to top of the table. All cooperative effects mentioned are positive. CT*

a

Homo- and heterotropic cooperativity mostly between Pi and G-I-P

Catalytic

Catalytic

Sites

T A B L E IV SUMMARY O F S I T E - S I T E INTERACTIONS IN PHOSPHORYLASE"

226

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

aspartate transcarbamylase in the presence of succinate and was explained by assuming that binding of the competitive inhibitor to one of the subunits of the enzyme would place the other subunits in the "active" con­ formation {50). It implies that each subunit of phosphorylase has a potentially operative catalytic site, but aggregation to dimer or tetramer is required to provide for the proper conformation. E. Interconversion of Phosphorylases b and a Interconversion of phosphorylases b and a is catalyzed by phosphorylase kinase and phosphatase, and when both processes occur simultaneously the overall reaction is that of an ATPase : 2 phosphorylase b + 4ATP

Mg 2+ kinase

► phosphorylase a + 4ADP

phosphatase

phosphorylase a + 4H 2 0 Sum:

4ATP + 4H 2 0

► phosphorylase b + 4Pi ► 4ADP + 4Pi

In the kinase reaction, the γ-phosphate of ATP is transferred to the protein; the reaction is essentially irreversible and no synthesis of ATP has been observed from 32P-labeled phosphorylase a and ADP under a variety of conditions (94). Calcium cannot replace Mg in the b to a reaction; on the contrary, it acts as a competitive inhibitor (the Km for Mg 2+ is 1.9 X 10"3 M vs. a K{ of 3.0 X IO"4 M for Ca 2+ as measured in purified systems) (92). However, since the concentration of free Mg is at least three orders of magnitude higher than that of Ca in the sarcoplasm, this competi­ tion cannot be of physiological significance. There are good reasons to believe that, at least in the case of rabbit muscle phosphorylase, neither phosphorylation nor dephosphorylation of the enzyme proceeds in an all-or-none fashion in which, at any one time, only a mixture of the fully phosphorylated a and fully dephosphorylated b species would be found (76). It was proposed that these reactions actually proceed in a stepwise manner in which partially phosphorylated inter­ mediates are produced according to the scheme depicted in Fig. 4 for the a —> b reaction. This hypothesis was based on the following set of observa­ tions (42) : The conversion of phosphorylase a —» b can be followed by the disappear­ ance of phosphorylase a activity (i.e., measured in the absence of AMP) or by loss of radioactivity if 32P-labeled phosphorylase a is used as substrate. Of course, one would expect the two rates to be similar, and this is indeed what is observed provided enzymatic activity is measured at low concentra­ tion of substrate (Km (glucose-1-P) for phosphorylase a is approximately 16 m l ) . However, if the reaction mixture is assayed at high concentration

227

CONTROL OF GLYCOGEN DEGRADATION

8^-18^8 · 8^88^88 1

11

11

11

11 11

t t 18 [18 88] 88 88 FIG. 4. Model for the formation of phospho-dephospho hybrids during the phosphorylase a to b conversion.

of glucose-1-P (approximately 75 m l ) , essentially no loss of activity is observed until approximately half the protein-bound phosphate has been hydrolyzed. Furthermore, addition of as little as 1 mikf glucose-6-P to this assay mixture totally suppresses this abnormally high initial activity, lowering it to a level identical if not below that of the bound phosphate. Analogous observations were made during the phosphorylase 6 —» a con­ version catalyzed by phosphorylase kinase : appearance of phosphorylase a activity runs ahead of 32P incorporation so that maximum activity is ob­ tained when only approximately half the 32P has been incorporated. Here again, this abnormally high activity is suppressed by inclusion of 1 m l glucose-6-P to the assay system. Since glucose-6-P has absolutely no effect on the activity of phosphorylase a (119), it is obvious that a new molecular entity having enzymatic properties different from those of the fully phos­ phorylated or dephosphorylated species must have been generated. To account for the large differences in enzymatic activities observed with and without glucose-6-P, the following assumption was made. Phosphoryl­ ase is known to exist in two conformations, active and inactive (depicted as squares and circles, respectively, in Fig. 5) in equilibrium with one another. In phosphorylase 6 this equilibrium strongly favors the inactive conformation unless a positive effector such as AMP is added; glucose-6-P counteracts this effect and favors the inactive state. Equilibrium for the phosphorylated form of the protein (depicted in black in Fig. 5) is strongly displaced toward the active conformation, to the extent that a totally inactive, phosphorylated species has never been seen under normal condi­ tions. By contrast, equilibrium between the active and inactive conforma­ tions of the partially phosphorylated hybrids would be much more sensitive toward positive or negative effectors, i.e., much more easily shifted from one state to the other. Perhaps, this simply results from the fact that phosphorylation of only one of the subunits introduces an element of dissym­ metry in the molecule that prevents it from settling down in the more stable

228

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

PHOSPHATASE

I -~Γ-Β Hi

66p

f-

66p

1-8-8

|[-

KINASE

FIG. 5. Model for the conformation of phosphorylases a and ò and phospho-dephospho hybrids. Circles represent inactive conformations, and squares, active ones. Open and filled symbols represent nonphosphorylated and phosphorylated subunits, respectively.

conformations of the parent molecules. In the presence of high concentra­ tions of glucose-1-P, the material would be fully active; on the other hand, glucose-6-P at concentrations of 10~3 M or below would displace the equilibrium entirely toward the inactive state. Because any active conformation of rabbit muscle phosphorylase tends to aggregate to a tetrameric state whereas the inactive conformation remains as a dimer, these shifts in structure can be readily visualized in the ultracentrifuge (42). Addition of glucose-1-P to a mixture of hybrids yields predominantly tetramers whereas addition of glucose-6-P produces mostly dimers; such effects are much less pronounced with a mixture of pure phosphorylase b and a. Of course, these are only gross manifestations of conformational changes; variations in the activity pattern of the hybrids might result from far more subtle alterations in tertiary structure or contact among the enzyme subunits that would require more sophisticated approaches to be visualized. A partial separation of hybrids was achieved by ion-exchange chromatography. Unfortunately, no pure hybrid preparation could be obtained as yet since the heterologous species rapidly dismutate to the thermodynamically more stable homologous forms of phosphorylase a and b (4®, 76). The existence of partially phosphorylated hybrids whose activity and susceptibility to positive and negative effectors is different from those of the b and a forms themselves would provide for a sophisticated way of regulating metabolism.

229

CONTROL OF GLYCOGEN DEGRADATION

III. State of Activity of Phosphorylase in Striated Muscle As indicated earlier, phosphorylase ò can be activated either by inter­ action with AMP or by covalent modification. The extent of the allosteric activation depends not only on the concentration of AMP but also, and to a large extent, on the conditions of the reaction, including nature and concentration of substrates and other effectors (particularly ATP and glucose-6-P), divalent metal ions, pH, temperature, etc. Whether or not this form of control is physiologically operative, and to what extent, has been argued from the moment the two forms of phosphorylase were dis­ covered. At first, it appeared from the early data of Lohmann and Schuster (99) that the concentration of AMP in resting muscle (approximately 0.01 m l ) was not high enough to appreciably activate phosphorylase b (22, 24)- With improvements in the techniques of "quick-freezing" of muscle tissue, the earlier values were revised upward to approximately 0.5 m l AMP, i.e., more than sufficient to maintain phosphorylase in an active state (to the extent that the physiological significance of the phos­ phorylase b —> a conversion was questioned) (49). The whole matter had to be reexamined when it was found that AMP activation was opposed by ATP (105, 123) and glucose-6-P and that the concentration of Pi in resting muscle was well below its Km value (119). In the following section, we shall attempt to discuss the current status of this problem. Table V summarizes some of the data regarding the cellular concentraTABLE V CONCENTRATIONS, Km, AND Ki OF SUBSTRATE AND EFFECTORS OF PHOSPHORYLASE IN RESTING MUSCLE 0

Substrate or effectors Pi Glucose-1-P Glycogen AMP ADP ATP Mg2+ Glucose-6-P

Concentration6 (ml)

Km (mM)

Ki (mM)

2 0.01 60 0.5 2 7 12 0.3

1-10 1.6-16 0.1 0.02-0.I e

— — —

— — —

3 3

—

0.3

° Values listed are gross averages and will vary with individual prepara­ tions and experimental conditions. 6 Given, per milliliter intracellular water, taken as 50% of tissue wet weight. c Depending on Pi concentration.

230

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

tions, Km or Ki of substrates and effectors of rabbit muscle phosphorylase. It should be emphasized that these values are merely averages: considerable fluctuations occur with the origin and physiological condition of the tissue under study and the experimental procedure used. Table VI lists the concentrations and ' 'potential activities" of some of the enzymes of the phosphorylase system and considers these values in relation to the breakdown of glycogen. It can be calculated that, in resting muscle, phosphorylase should display approximately 1-2% of its potential activity, if the compounds listed in Table V were the sole factors involved. This value has indeed been found in a reconstituted system in which purified phosphorylase b was placed under conditions approximating those listed in Table V, and corresponded to a rate of production of glucose-1-P of 1-2 ^moles/min/ml* (119) (see also 15). Yet, it has been shown that production of lactic acid in resting muscle is of the order of 10-30 nmole/min/ml (69) or approximately 100 times less than expected. Similar values for glycolysis were obtained when C0 2 production was measured assuming that it originated exclusively from glucose oxidation. As will be seen below, the rate-limiting step in glycolysis does appear to be that catalyzed by glycogen phosphorylase. This indicates that in resting muscle, phosphorylase must be essentially inactive, i.e., have an activity of the order of 1/10,000 its total potential activity (Table VI). There are several further indications that AMP activation of phosphoryl­ ase cannot play a major role in determining the breakdown of glycogen (15, 69). Helmreich and Cori (69) have shown that changes in the level of nucleotide effectors in the muscle are not fast enough to account for the rapid (10-20 seconds at most) activation of phosphorylase observed upon electric stimulation: whereas the level of creatine-P falls rapidly, levels of ATP, ADP, glucose-6-P, and in particular AMP change hardly at all during the first 5-10 minutes following the onset of contraction and cer­ tainly not nearly as much as would be required for an activation of phos­ phorylase (188, 139). Of course, conclusions such as these are based on the assumption that within the muscle cell (a) there is no compartmentation where AMP could preferentially accumulate, and (b) that phosphorylase would have enzymatic characteristics identical to those measured on the pure enzyme. The latter is a dangerous assumption since muscle phosphoryl­ ase, together with several other enzymes involved in glycogen metabolism, appears to be associated with a glycogen particulate fraction (see Sec­ tion VI). * All concentrations are given per milliliter of intracellular water, the latter taken as half the weight of the fresh tissue {120).

TABLE VI

(0.3) A 4.7

— 4 SS interchanges, only three types of chemical substitutions have been de­ scribed, namely, phosphorylation, adenylylation, and ADP-ribosylation. New examples have appeared recently; no doubt others will soon follow. There remains of course the fundamental question as to how covalent modification brings about those particular changes in conformation that result in the appearance or disappearance of enzymatic activity (and loss of cooperativity in the case of phosphorylase). Questions such as these will have to await the elucidation of the primary and tertiary structure of regulatory enzymes by means of sequence analysis and X-ray crystal­ lography. In addition to describing the control of enzymes by allosteric and co-

246

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

valent modifications, this review called attention to regulatory mechanisms that rely on protein-protein—or other kinds of macromolecular—inter­ action. No doubt additional examples will come to light with a better understanding of intracellular organization. Such multicomponent systems, in addition to allowing for specific interactions among control enzymes and their respective protein substrates, could provide for a sort of "chemical compartmentation" in which the properties of the enzymes involved could be considerably altered. The phosphorylase system is regulated by a highly complex mechanism in which several enzymes act on one another. It has been proposed here that the main reason for this multiplicity of steps is to allow for a con­ certed regulation of several physiological processes. Do such metabolic interrelationships exist in other systems? Many other questions remain to be answered. For instance, it is well known that adenine nucleotides play an important role in the regulation of certain pathways, particularly those concerned with energy metabolism. Do the many enzymes that are affected by interaction with this class of compounds have common structural features? For sure, adenine nucleotides would be expected to bind readily to many proteins by virtue of their hydrophilic character, multiplicity of charges, and polyheterocyclic nature. Do many types of interactions occur though the end result is always similar, or do these enzymes have analogous sites that perhaps all evolved from a common, ancestral, nucleotide-binding entity? In the case of glycogen degradation and its relation to muscle contraction, one wonders how the two regulatory mechanisms operate, i.e., simul­ taneously or in sequence? For instance, is covalent control operative mainly at the onset of contraction while AMP activation takes over during the recovery period? In view of the multiple bands displayed by kinase upon disc gel electrophoresis in sodium dodecyl sulfate the enzyme might possess different types of subunits. Since conformational changes in both kinase and troponin are triggered by essentially the same concentrations of Ca 2+ ions, it would be tempting to speculate that kinase might possess a troponin-like subunit which, in the absence of Ca 2+ ions, represses its activity. Could there be other structural relationships between proteins involved in contraction and metabolism? What is the mechanism by which Ca is taken up and released by elements of the sarcoplasmic reticulum; do the latter contain specific calcium-binding entities? Finally, since there appears to exist several protein kinases all activated by cyclic-AMP, what determines which should become operative at any one time? The study of the phosphorylase system has added much to our knowledge of the molecular basis of hormonal action (12, 56, 134) and the control of cellular processes. When considered together with the remarkable advances

CONTROL OF GLYCOGEN DEGRADATION

247

made in the field of muscle contraction (35, 36, 79, 125), one can hope that the complex interrelations existing between these physiological processes will soon be understood. ACKNOWLEDGMENTS

The authors are much indebted to Professor Ernst Helmreich and Dr. Philip Cohen for their help and many fruitful suggestions in the preparation of this manuscript. REFERENCES

1. Abdullah, M., Fischer, E. H., Qureshi, M. Y., Slessor, K. N., and Whelan, W. J., Biochem. J. 97, 9P (1965). 2. Appleman, M. M., Yunis, A. A., Krebs, E. G., and Fischer, E. H., J. Biól. Chem. 238, 1358 (1963). 3. Atkinson, D. E., Annu. Rev. Biochem. 35, 611 (1966). 4. Avramovic-Zikic, O., Smillie, L. B., and Madsen, N. B., / . Biol. Chem. 245, 1558 (1970). 5. Battell, M. L., Smillie, L. B., and Madsen, N. B., Can. J. Biochem. 46, 609 (1968). 6. Battell, M. L., Zarkadas, C. G., Smillie, L. B., and Madsen, N. B., J. Biol. Chem. 243,6202 (1968). 7. Boeker, E. A., Fischer, E. H., and Snell, E. E., J. Biol. Chem. 246, in press (1971). 8. Bresler, S., and Firsov, L., / . Mol. Biol. 35, 131 (1968). 9. Brostrom, C. O., Hunkeler, F. L., and Krebs, E. G., J. Biol. Chem. 246, 1961 (1971). 10. Brown, D. H., and Cori, C. F., in "The Enzymes" (P. D. Boyer, H. Lardy, and K. Myrbäck, eds.), 2nd ed., Vol. 5, p. 207. Academic Press, New York, 1961. 11. Bue, M. H., and Bue, H., Broc. J,th Meeting Fed. Eur. Biochem. Soc, Oslo, 1967. 12. Butcher, R. W., Robison, G. A., Hardman, J. G., and Sutherland, E. W., Advan. Enzyme Regul. 6, 357 (1968). 13. Chelala, C. A., Hirschbein, L., and Torres, H. N., Proc. Nat. Acad. Sci. U. S. 68, 152 (1971). 13a. Chelala, C. A., and Torres, H. N., Biochim. Biophys. Acta 178, 423 (1969). 14. Chignell, D. A., Gratzer, W. B., and Valentine, R. C , Biochemistry 7, 1082 (1968). 15. Childress, C. C , and Sacktor, B., / . Biol. Chem. 245, 2927 (1970). 16. Childress, C. C , Sacktor, B., Grossman, I., and Bueding, E., J. Cell Biol. 45, 83 (1970). 17. Cohen, P., Duewer, T., and Fischer, E. H., Biochemistry 10, 2683 (1971). 18. Cohn, M., / . Biol. Chem. 180, 771 (1949). 19. Cohn, M., and Cori, G. T., J. Biol. Chem. 175, 89 (1948). 20. Corbin, J. D., Reimann, E. M., Walsh, D. A., and Krebs, E. G., J. Biol. Chem. 245,4849 (1970). 21. Cori, C. F., Physiol. Rev. 11, 143 (1931). 22. Cori, C. F., in "Enzymes: Units of Biological Structure and Function" (O. H. Gaebler, ed.), p. 573. Academic Press, New York, 1956. 23. Cori, C. F., Cori, G. T., and Green, A. A., J. Biol. Chem. 151, 39 (1943). 24. Cori, G. T., / . Biol. Chem. 158, 333 (1945). 25. Cori, G. T., and Cori, C. F., / . Biol. Chem. 135, 733 (1940). 26. Cori, G. T., and Cori, C. F., / . Biol. Chem. 158, 321 (1945). 27. Cortijo, M., and Shaltiel, S., Biochem. Biophys. Res. Commun. 39, 212 (1970).

248

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

28. 29. 30. 31.

Cortijo, M., Steinber, L. Z., and Shaltiel, S., J. Biol. Chem. 246, 933 (1971). Cowgill, R. W., / . Biol. Chem. 234, 3146 (1959). Damjanovich, S., and Kleppe, K., Biochim. Biophys. Acta 122, 145 (1966). Danforth, W. H., Helmreich, E., and Cori, C. F., Proc. Nat. Acad. Sci. U. S. 48, 1191 (1962). Danforth, W. H., and Lyon, J. B., J. Biol. Chem. 239, 4047 (1964). DeLange, R. J., Kemp, R. G., Riley, R. D., Cooper, R. A., and Krebs, E. G., / . Biol. Chem. 243, 2200 (1968). DeVincenzi, D . L., and Hedrick, J. L., Biochemistry 6, 3489 (1967). Ebashi, S., and Endo, M., Progr. Biophys. Mol. Biol. 18, 123 (1968). Ebashi, S., Endo, M., and Ohtsuki, I., Quart. Rev. Biophys. 2, 351 (1969). Ebashi, S., and Lipmann, F., / . Cell Biol. 14, 389 (1962). Feldman, K., and Helmreich, E., personal communication. Fischer, E. H., Struct. Activ. Enzymes, Fed. Eur. Biochem. Soc. Symp. 1st, 1964· Fischer, E . H., Forrey, A. W., Hedrick, J. L., Hughes, R. C , Kent, A. B., and Krebs, E. G., in "Chemical and Biological Aspects of Pyridoxal Catalysis" (E. E . Snell, P. M. Fasella, A. Braunstein, and A. Rossi-Fanelli, eds.), p. 543, Pergamon, Oxford, 1963. Fischer, E . H., Graves, D . J., Crittenden, E. R., and Krebs, E . G., J. Biol. Chem. 234, 1698 (1959). Fischer, E . H., Hurd, S. S., Koh, P., Seery, V. L., and Teller, D . C , Fed. Eur. Biochem. Soc, 1968, p . 19. Fischer, E. H., Kent, A. B., Sneider, E. R., and Krebs, E. G., J. Amer. Chem. Soc. 80,2906 (1958). Fischer, E . H., and Krebs, E . G., J. Biol. Chem. 216, 121 (1955). Fischer, E . H., and Krebs, E. G., Fed. Proc. 25, 1511 (1966). Fischer, E . H., Pocker, A., and Saari, J. C , in "Essays in Bio chemistry' ' (P. N . Campbell and F . Dickens, eds.), p . 23. Academic Press, New York, 1970. Forrey, A. W., Sevilla, C. L., Saari, J. C , and Fischer, E . H., Biochemistry 10, 3132 (1971). Fosset, M., Nielsen, L. D., Muir, L. W., and Fischer, E. H., in preparation. Gerbach, E., Deuticke, B., and Dreisbach, R. H., Naturwissenschaften 50, 228 (1963). Gerhart, J. C , and Pardee, A. B., / . Biol. Chem. 237, 891 (1962). Goff, C. G., and Weber, K., Cold Spring Harbor Symp. Quant. Biol. 35, 101 (1970). Gold, A. M., Biochemistry 7, 2106 (1968). Gold, A. M., and Osber, M. P., Biochem. Biophys. Res. Commun. 42, 469 (1971). Graves, D . J., Fischer, E. H., and Krebs, E. G., / . Biol. Chem. 235, 805 (1960). Graves, D . J., Scharfenberg-Mann, S. A., Philip, G., and Oliveira, R. J., J. Biol. Chem. 243, 6090 (1968). Greengard, P., and Costa, E., eds., "Role of Cyclic A M P in Cell Function." Raven, New York, 1970. Guirard, B. M., and Snell, E. E., in "Comprehensive Biochemistry" (M. Florkin and E. H. Stotz, eds.), Vol. 15, p. 138. Elsevier, Amsterdam, 1964. Hansford, R. G., and Sacktor, B., FEBS Lett. 7, 183 (1970). Haschke, R. H., unpublished results. Haschke, R. H., Meyer, F., Heilmeyer, L., Jr., and Fischer, E. H., J. Biol. Chem. 245,6657 (1970). Hedrick, J. L., and Fischer, E. H., Biochemistry 4, 1337 (1965). Hedrick, J. L., Shaltiel, S., and Fischer, E . H., Biochemistry 6, 2117 (1966). Hedrick, J. L., Shaltiel, S., and Fischer, E. H., Biochemistry 8, 2422 (1969).

32. 33. 34. 35. 36. 37. 38. 39. 40.

4L 42. 43. 44· 45. 46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63.

CONTROL OF GLYCOGEN DEGRADATION

249

64. Heilmeyer, L., Jr., Meyer, F., Haschke, R. H., and Fischer, E. H., J. Biol. Chem. 245,6649 (1970). 65. Helmreich, E., personal communication. 66. Helmreich, E., FEBS Symp. 19, 131 (1969). 67. Helmreich, E., in ''Comprehensive Biochemistry" (M. Florkin and E. H. Stotz, eds.), Vol. 17, p. 17. Elsevier, Amsterdam, 1969. 68. Helmreich, E., and Cori, C. F., Proc. Nat. Acad. Sci. U. S. 61, 131 (1964). 69. Helmreich, E., and Cori, C. F., Advan. Enzyme Regul. 3, 91 (1965). 70. Hers, H. G., DeWulf, H., and Stalmans, W., FEBS Lett. 12, 73 (1970). 71. Hohorst, H. J., Reim, M., Bartels, H., Biochem. Biophys. Res. Commun. 7, 137 (1962). 72. Horjo, T., Nishizuma, Y., Hayaishi, O., and Kato, I., / . Biol. Chem. 243, 3553 (1968). 73. Hughes, R. C , Yunis, A. A., Krebs, E. G., and Fischer, E. H., / . Biol. Chem. 237, 40 (1962). 74. Hullar, T. L., Tetrahedron Lett. 49, 4921 (1967). 75. Hurd, S. S., Ph.D. Thesis, University of Washington, 1967. 76. Hurd, S. S., Teller, D. C, and Fischer, E. H., Biochem. Biophys. Res. Commun. 24, 79 (1966). 77. Huston, R. B., and Krebs, E. G., Biochemistry 7, 2116 (1968). 78. Huttunen, J. K., Steinberg, D., and Mayer, S. E., Proc. Nat. Acad. Sci. U. S. 67, 290 (1970). 79. Huxley, H. E., Science 164, 1356 (1969). 80. Illingworth, B., Jansz, H. S., Brown, D. H., and Cori, C. F., Proc. Nat. Acad. Sci. U. S. 44, 1180 (1958). 80a. Johnson, J. F., Tu, J.-L, Shonka Bartlett, M. L., and Graves, D. J., J. Biol. Chem. 246, 5560 (1970). 80b. Kahn, V., and Blum, J. J., Arch. Biochem. Biophys. 143, 80 (1971). 81. Karpatkin, S., Helmreich, E., and Cori, C. F., J. Biol. Chem. 239, 3139 (1964). 82. Kastenschmidt, L. L., Kastenschmidt, J., and Helmreich, E., Biochemistry 7, 3590 (1968). 83. Kastenschmidt, L. L., Kastenschmidt, J., and Helmreich, E., Biochemistry 7, 4543 (1968). 84. Keller, P. J., and Cori, G. T., Biochim. Biophys. Acta 12, 235 (1953). 85. Keller, P. J., and Cori, G. T., / . Biol. Chem. 214, 127 (1955). 86. Kent, A. B., Krebs, E. G., and Fischer, E. H., J. Biol. Chem. 232, 549 (1958). 87. Kingdon, H. S., Shapiro, B. M., and Stadtman, E. R., Proc. Nat. Acad. Sci. U. S. 68, 1703 (1967). 88. Krebs, E. G., personal communication. 89. Krebs, E. G., and Fischer, E. H., Biochim. Biophys. Acta 20, 150 (1956). 90. Krebs, E. G., and Fischer, E. H., Advan. Enzymol. 24, 263 (1962). 91. Krebs, E. G., and Fischer, E. H., Vitam. Horm. (New York) 22, 399 (1964). 92. Krebs, E. G., Graves, D. J., and Fischer, E. H., J. Biol. Chem. 234, 2867 (1959). 93. Krebs, E. G., Huston, R. B., and Hunkeler, F. L., Advan. Enzyme Regul. 6, 245 (1969). 94. Krebs, E. G., Kent, A. B., and Fischer, E. H., J. Biol. Chem. 231, 73 (1958). 95. Krebs, E. G., Love, D. S., Bratvold, G. E., Trayser, K. A., Meyer, W. L., and Fischer, E. H., Biochemistry 3, 1022 (1964). 96. Kumon, A., Yamamura, H., and Nishizuka, Y., Biochem. Biophys. Res. Commun. 41, 1290 (1970). 97. Leloir, L. F., Contr. Glycogen Metal·., Ctàa Found. Symp., 1963, p. 68 (1964).

250

E. H. FISCHER, L. M. G. HEILMEYER, JR., AND R. H. HASCHKE

98. Liebig, J., in "Aus Justus Liebig's und Friedrich Wöhler's Briefwechsel in den Jahren 1829-1873" (A. W. Hofmann, ed.), Vol. II. Braunschweig, 1888. 99. Lohmann, K., and Schuster, P., Biochem. Z. 272, 24 (1934). 100. Lowry, H. 0., Passonneau, J. V., Hasselberger, F. X., and Schulz, D. W., J. Biol. Chem. 239, 18 (1964). 101. Lynen, F., Fed. Proc. 20, 941 (1961). 102. Lyon, J. B., and Porter, J., J. Biol. Chem. 238, 1 (1963). 103. Lyon, J. B., Porter, J., and Robertson, M., Science 155, 1550 (1967). 104. McClintock, D. K., and Markus, G., J. Biol. Chem. 243, 2855 (1968). 105. Madsen, N. B., Biochem. Biophys. Res. Commun. 6, 310 (1961). 106. Madsen, N. B., Biochem. Biophys. Res. Commun. 15, 390 (1964). 107. Madsen, N. B., and Cori, C. F., / . Biol. Chem. 223, 1055 (1956). 108. Madsen, N. B., and Shechosky, S., / . Biol. Chem. 242, 3301 (1967). 109. Martello, O. J., Woo, S. L., Riemann, E. M., and Davie, E. W., Bioehemistry 9,4807 (1970). 110. Merlevede, W., and Riley, G. A., / . Biol. Chem. 241, 3517 (1966). 111. Metzger, B. E., Glaser, L., and Helmreich, E., Bioehemistry 7, 2021 (1968). 112. Metzger, B. E., Helmreich, E., and Glaser, L., Proc. Nat. Acad. Sci. U. S. 57,994 (1967). 113. Meyer, F., Heilmeyer, L., Jr., Haschke, R. H., and Fischer, E. H., / . Biol. Chem. 245,6642 (1970). 114. Meyer, W. L., Fischer, E. H., and Krebs, E. G., Bioehemistry 3, 1033 (1964). 115. Meyerhof, O., "Die chemischen Vorgänge im Muskel," Monogr. Physiol., Vol. 22. Berlin, 1930. 116. Milroy, T. H., Physiol. Rev. 11, 515 (1931). 117. Mommaerts, W. F. H. M., Illingworth, B., Pearson, C. M., Guillory, R. J., and Serayderian, K., Proc. Nat. Acad. Sei. U. S. 45, 791 (1959). 118. Monod, J., Changeux, J. P., and Jacob, F., J. Mol. Biol. 6, 306 (1963). 119. Morgan, H. E., and Parmeggiani, A., / . Biol. Chem. 239, 2440 (1964). 120. Narahara, H. T., Özand, P., and Cori, C. F., / . Biol. Chem. 235, 3370 (1960). 121. Nolan, C , Novoa, W. B., Krebs, E. G., and Fischer, E. H., Bioehemistry 3, 542 (1966). 122. Ozawa, E., Hosoi, K., and Ebashi, S., J. Biochem. {Tokyo) 61, 531 (1967). 123. Parmeggiani, A., and Morgan, H. E., Biochem. Biophys. Res. Commun. 9, 252 (1962). 124. Parnass, J. K., Annu. Rev. Biochem. 1, 431 (1932). 125. Peachey, L. D., Annu. Rev. Phys. 30, 401 (1968). 126. Pecararo, R. E., M. S. Thesis, University of Washington (1969). 127. Pocker, A., and Fischer, E. H., Bioehemistry 8, 5181 (1969). 128. Portzell, H., Caldwell, P. C , and Rüegg, J. C , Biochim. Biophys. Ada 79, 581 (1964). 129. Posener, J. B., Stern, R., and Krebs, E. G., / . Biol. Chem. 240, 982 (1965). 129a. Puchwein, G., Kratky, O., Gölker, C. F., and Helmreich, R., Bioehemistry 9, 4691 (1970). 130. Rändle, P. G., Symp. Soc. Exp. Biol. 18, 129 (1964). 131. Reed, L. J., Curr. Top. Cell Regul. 1, 233 (1969). 132. Reimann, E. M., Brostrom, C. O., Corbin, J. D., King, C. A., and Krebs, E. G., Biochem. Biophys. Res. Commun. 42, 187 (1971). 132a. Reimann, E. M., Walsh, D. A., and Krebs, E. G., J. Biol. Chem. 246,1986 (1971). 133. Riley, R. D., DeLange, R. J., Bratvold, G. E., and Krebs, E. G., J. Biol. Chem. 243,2209 (1968).

CONTROL OF GLYCOGEN DEGRADATION

251

I34. Robison, G. A., Butcher, R. W., and Sutherland, E. W., Annu. Rev. Biochem. 37, 149 (1968). 185. Rollston, F. S., and Newsholme, E. A., Biochem. J. 104, 524 (1967). 136. Saari, J. C., Ph.D. Thesis, University of Washington (1970). 137. Sabo, D., unpublished results. 138. Sacktor, B., and Hurlbut, E. C , J. Biol. Chem. 241, 632 (1966). 139. Sacktor, B., and Wormser-Shavit, E., / . Biol. Chem. 241, 624 (1966). 140. Schmid, R., and Mahler, R., J. Clin. Invest. 38, 2044 (1959). 141. Schwartz, M., and Hofnung, M., Eur. J. Biochem. 2, 132 (1967). 142. Scrutton, M. C , and Utter, M. F., Annu. Rev. Biochem. 37, 249 (1968). 143. Seery, V. L., Fischer, E. H., and Teller, D. C , Biochemistry 6, 3315 (1967). 144. Seery, V. L., Fischer, E. H., and Teller, D. C , Biochemistry 9, 3591 (1970). 145. Sevilla, C. L., Ph.D. Thesis, University of Washington (1969). 146. Sevilla, C. L., and Fischer, E. H., Biochemistry 7, 2161 (1968). 147. Shaltiel, S., Hedrick, J. L., and Fischer, E. H. Biochemistry 5, 2108 (1966). 148. Shaltiel S. Hedrick, J. L., and Fischer, E. H., Biochemistry 8, 2429 (1969). 149. Shaltiel, S., Hedrick, J. L., Pocker, A., and Fischer, E. H., Biochemistry 8,5189 (1969). 150. Soderling, T. R., Hickenbottom, J. P., Reimann, E. M., Hunkeler, F. L., Walsh, D. A., and Krebs, E. G., / . Biol. Chem. 245, 6317 (1970). 151. Strausbauch, P. H., and Fischer, E. H., Biochemistry 9, 233 (1970). 152. Sutherland, E. W., in "Phosphorus Metabolism" (W. D. McElroy and B. Glass, eds.), Vol. 1, p. 53. The Johns Hopkins Press, Baltimore, Maryland, 1941. 153. Takeda, M., Yamamura, H., and Ohga, Y., Biochem. Biophys. Res. Commun. 42, 103 (1971). 154. Tao, M., Salas, M. L., and Lipmann, F., Proc. Nat. Acad. Sci. U. S. 67, 408 (1970). 154a. Tracy, L. C , Pettit, F. H., and Reed, L. J., Proc. Nat. Acad. Sci. U. S. 62, 234 (1969). 155. Tu, J.-L, Jacobson, G. R., and Graves, D. J., Biochemistry 10, 1229 (1971). 156. Ullmann, A., Goldberg, M. E., Perrin, D., and Monod, J., Biochemistry 7, 261 (1968). 157. Ullmann, A., Vagelos, P. R., and Monod, J., Biochem. Biophys. Res. Commun. 17, 86 (1964). 158. Underwood, A. H., and Newsholme, E. A., Biochem. J. 104, 300 (1967). 159. Valentine, R. C., and Chignell, D. A., Nature (London) 218, 950 (1968). 160. Vidgoff, J., Ph.D. Thesis, University of Washington (1970). 161. Villar-Palasi, C., and Larner, J., Arch. Biochem. Biophys. 94, 436 (1961). 162. Voet, J. G., and Abeles, R. H., J. Biol. Chem. 245, 1020 (1970). 162a. Walsh, D. A., Ashby, C. D., Gonzalez, C , Calkins, D., Fischer, E. H., and Krebs, E. G., / . Biol. Chem. 246, 1977 (1971). 162b. Walsh, D. A., Perkins, J. P., Brostrom, C. O., Ho, E. S., and Krebs, E. G., J. Biol. Chem.24Q, 1968 (1971). 163. Walsh, D. A., Perkins, J. P., and Krebs, E. G., / . Biol. Chem. 243, 3763 (1968). 164. Wang, J. H., and Graves, D. J., Biochemistry 3, 1437 (1964). 165. Wang, J. H., and Tu, J.-L, J. Biol. Chem. 246, 176 (1970). 166. Wanson, J. C., and Drochmans, P., J. Cell Biol. 38, 130 (1968). 167. Wieland, O., and Siess, E., Proc. Nat. Acad. Sci. U. S. 65, 947 (1970). 168. Wolf, D. P., Fischer, E. H., and Krebs, E. G., Biochemistry 9, 1923 (1970). 169. Wu, R., J. Biol. Chem. 240, 2373 (1965). 170. Wulff, K., Mecke, D., and Holzer, H., Biochem. Biophys. Res. Commun. 28, 740 (1967). 171. Yunis, A. A., and Krebs, E. G., J. Biol. Chem. 237, 34 (1962).

Author Index Numbers in parentheses are reference numbers and indicate that an author's work is referred to, although his name is not cited in the text. Numbers in italics show the page on which the complete reference is listed.

Abdullah, M., 223(1), 247 Abeles, R. H., 223(162), 251 Abraham, S., 123(1), 141(141), 158(2), 162, 165 Abramovitz, A. S., 139(108), 157(27, 108, 116), 158(108, 116), 162, 164, 165 Ackers, G. K., 81(17), 114 Adelberg, E . A., 21(11), 37 Ahmad, F., 132(3, 64), 162, 168 Ailhaud, G. P., 141(163), 142(163), 143(4, 5, 44, 163), 162, 163, 166 Alberts, A. W., 123, 126(160, 161), 127(9), 129(8), 131(107), 132(107), 133(161), 135(8,9), 141(163), 142(92, 163), 143 (163), 144(93, 94), 146(6), 147(7, 51), 148(51), 154(7), 162, 163, 164, 166 Alberty, R. A., 56, 72 Allman, D . W., 139, 159(10, 40), 162, 163 Anderson, B . M., 99(82), 115 Anderson, C. D., 99(82), 115 Anderson, P . J., 83(33), 114 Anfinsen, C. B., 85(40), 87(40), 101(40), 115 Anthony, D . D., 43(2), 72 Antonini, E., 177, 178(3, 8), 180(2, 8), 181(2), 182,183(1, 2, 4), 192(1), 206, 207 Appella, E., 79, 114 Appleman, M . M., 247 Arditti, R. R., 66(27), 68(27), 73 Ashby, C. D., 234(162a), 251 Atkinson, D . E., 168, 207, 243(3), 247 Avramovic-Zikic, O., 216(4), 247

B Baker, J. P., 99(88), 116 Bakerman, H . A., 152(76), 164 Ball, E . G., 120(37, 182), 163, 166 Ballard, F . J., 120(11), 162 Balthasar, N., 176(101), 209 Bambers, G., 102(100), 116 253

Banaszak, L. J., 171(114), 184(114), 210 Barber, E . D., 202, 207 Barbour, S. D., 41(3), 72 Bard, J. R., 83, 89(59), 114, 115 Barker, H. A., 84(38), 102(38), 114 Barnes, E . M., Jr., 142, 158(12), 162 Barratt, R. W., 84, 114 Bartels, H., 232(71), 249 Bartley, W., 103(103), 105(103), 116 Bates, D . J., 86(45), 90(60), 115 Battell, M . L., 216(5, 6), 247 Bauer, H., 150(114), 165 Baum, H., 159(23), 161(23), 162 Bautz, E . K. F., 43(14), 45(14), 73 Bayley, P . M., 83, 114 Béchet, J., 3(3, 34), 4(4), 5(4), 6(1, 2, 4, 26), 14(26), 21(3, 34, 40), 23(34), 24(3, 34, 40), 37(4), 87, 38 Beckwith, J. R., 39(5), 40(5,41), 41(4), 43(105), 63(89), 66(27, 90,105), 67(35, 77), 68(27), 72, 73, 74, 75 Béguin, S., 60(40), 73 Bell, R., 103(102), 104(102), 116 Benesch, R., 174(7), 178(7), 207 Benesch, R. E., 174(7), 178(7), 207 Bennett, N . G., 204(50), 208 Berg, P., 43(39, 104), 47(39), 73, 75 Berger, R. L., 178(8), 180(8), 207 Bernhardt, W., 103(104), 105(104), 116 Betheil, J. J., 185(79), 209 Beyreuther, K., 43(58), 44(6, 58), 48(6), 62(6), 72, 74 Bhaduri, A., 120(144), 165 Birge, C. H., 149(52, 53), 163 Birnbaum, J., 140(13, 14), 162 Bitensky, M . W., 79, 101(6, 93, 94), 114, 116 Bittman, R., 185(73), 187(73), 197(73), 208 Bloch, K., 141(19), 148(15, 77), 159(60), 162, 168, 164 Bloch, W. A., 185(9), 186(9), 207

254 Blum, J. J., 224(80b), 249 Bock, R. M., 156(185), 157(24, 184, 185), 162, 166 Boeker, E. A., 218(7), 247 Boezi, J. A., 60(7), 72, 102(97), 104(97), 116 Bone, D. H., 110(123), 116 Borst, P., 103(107), 105(107), 107(107), 111, 116, 117 Bortz, W. M., 123, 133(109), 135(16, 17, 109), 136(109), 162, 165 Bourgeois, C. M., 23, 37 Bourgeois, S., 14(6), 37, 41(9), 42(8, 78, 80, 81, 82, 84), 43(78, 81), 44(8, 10, 12, 78, 84), 46(78, 84), 47(10, 78, 84), 48 (10), 49(8, 38, 82), 50(78, 82, 84), 51 (80, 81, 84), 52(12, 79, 84), 53(82), 54 (80), 55(80, 84), 56(80), 57(82), 60(11, 80), 62(10, 38), 63(12, 79), 66(82), 72, 73, 74 Brächet, P., 70(24), 71(24), 73 Bradley, R. M., 160(128), 165 Bradshaw, R. A., 83(31), 99(86), 100(86),

114, ne

Brady, R. O., 123(18), 160(128), 162, 165 Brand, L., 171(48), 208 Branscomb, E. W., 60(13), 73 Brattin, W. J., 79(5), 80(5), 100(5), 114 Bratvold, G. E., 233(95), 235(133), 249, 250 Bresler, S., 219(8), 247 Bright, H. J., 202, 207 Brindley, D. N., 141(19),^^ Brosnan, J. T., 112(134), 117 Brostrom, C. O., 233(9), 234(9, 162b), 235 (132), 238(9), 240(9), 243(162b), 247, 250 Brown, N. C , 170(10), 174(10), 207 Brown, T. H., 212(10), 216(80), 219(80), 247, 249 Brunori, M., 177, 178(3, 8), 180(2, 8, 11), 183(1, 2, 4), 192(1), 206, 207 Bulen, W. A., 110(124), 117 Bue, H., 174(93), 206(93), 209, 214(11), 247 Bue, M. H., 214(11), 247 Bucker, N. L. R., 123(20), 159(20), 162 Buckman, T., 194(12), 207 Bueding, E., 238(16), 247 Burchard, W., 79, 114

AUTHOR INDEX

Burgess, R. R., 43(14), 45(14), 73 Burstein, C , 53, 73 Burton, D. N., 141(22), 158(22), 161(21), 162 Bussel, J. B., 194(82), 209 Butcher, R. W., 234(12,134), 246(12,134), 247, 251 Butow, R. A., 98(80), 115 Butterworth, P. H. W., 156(185), 157(24, 27, 28,62,185), 158(117), 159(23), 161 (23), 162, 163, 165, 166 C Caldwell, P. C , 238(128), 250 Calkins, D., 234(162a), 251 Cano, A., 103(106), 105(106), 116 Capecchi, M. R., 41(16), 73 Carey, E. M., 158(25), 162 Cashel, M., 70(94), 75 Cassman, M., 81 (18a), 114 Caughey, W. S., 86, 115 Chadwick, P., 43(17), 44(71), 70(17, 71), 73, 74 Chaikoff, I. L., 123(1), 158(2), 162 Chamberlin, M. J., 194(118), 210 Chambers, D. A., 66(18), 73 Chamness, G. C , 49(19), 73 Chance, B., 173(13), 191(14), 207 Chang, H. C , 123(49), 124(49), 126(49), 133(131), 134(131), 135(131), 159(26), 162, 163, 165 Changeux, J. P., 3(31), 18(7, 30), 37, 38, 47(53, 55), 61(55), 74, 95(67), 115, 223 (118), 250 Changeux, J.-P., 167(85), 168(85, 96), 169 (85, 96), 171(85, 96), 182(85, 96), 185 (85, 96), 188(85, 96), 191(85), 192(85, 96), 194(85), 195(15), 197(17, 85), 198 (85, 96), 201(85, 96), 202(85), 205(85), 206(85), 207, 209 Chappelet, D., 204(77a), 209 Chelala, C. A., 241 (13a), 247 Chen, B., 43(68), 66(22), 67(22), 73, 74 Chesterton, C. J., 157(27, 28), 158(117), 162, 165 Chiancone, E., 178(3), 206 Chignell, D. A., 213(159), 214(14), 247, 251 Childress, C. C , 224(15), 230(15), 238(16), 247

255

AUTHOR INDEX

Chun, P. W., 81(17), II4 Churchich, J. E., 99(82), 116 Ciotti, M. M., 85(39), 115 Clark, A. J., 21(11), 37 Cleland, W. W., 170(16), 207 Coats, J. H., 20(8), 37 Coche, M., 18(15), 37 Coddington, A., 105(111), 107(111), 116 Coffee, C. J., 83(31), 99(86), 100(86), 114, 116 Cohen, G. N., 170(17), 199(17, 107), 200 (17, 67, 107, 109, 110), 201(18, 65, 67, 108, 109), 203(67), 207, 208, 209 Cohen, L. A., 175, 205(19), 207 Cohen, P., 213(17), 214(17), 222(17), 247 Cohen, P. P., 84(34), 87(34, 48), 97(71), 114, 115 Cohen-Bazire, G., 53(54), 74 Cohn, M., 41(9, 102), 42(80, 81), 43(81), 44(20, 88), 51(80, 81), 53(15, 54), 54(80), 55(80), 56(80), 60(80), 61(102), 72, 73, 74, 75, 223(18, 19), 247 Collins, J. M., 161(21), 102 Collins, K. D., 195(20, 21), 207 Colman, R. F., 79, 93(65), 94, 98(66), 99 (65), 114, 115 Colowick, S. P., 184(22), 207 Cooper, R. A., 232(33), 233(33), 234(33) 243(33), 248 Connaway, S., 66(27), 68(27), 73 Conway, A., 185(23), 186(23), 207 Cook, R. A., 185, 207 Corbin, J. D., 235(132), 243(20), 247, 250 Cori, C. F., 212(10, 21), 213(26), 216(80), 219(80), 222(23, 25), 223(81), 224(68), 229(22), 230(69), 240(31), 247, 248, 249, 250 Cori, G. T., 8(9), 37, 213(26, 84, 85), 222 (23, 25), 223(19), 229(24), 237(84), 247, 249 Corman, L., 84, 87(49), 88,97(49), II4,115 Cornish-Bowden, A., 168(25), 169(25), 207 Cortijo, M., 217(27, 28), 219(27, 28), 247 248 Costa, E., 246(56), 248 Cowgill , R . W. , 222(29) , 248 Cowie, D. E., 60(7), 72 Crapo, L., 41(59), 43(59), 74 Criddle, R. S., 143(139), 165 Crittenden, E. R., 222(41), 248

Cross, D. G., 82(20, 21, 22), 87(52), 93 (52), 98(75), II4, 115 Crothers, D. M., 51, 74 Cullen, J., 148(70), 163 Curran, J., 97(72, 74), 115

D Dahlen, J. V., 159(29), 161(29), 162 Daikuhara, Y., 120(148), 166 Dakshinamurti, K., 139(30, 31), 162 Dalziel, K., 87, 88, 115, 179(25a), 207 Damjanovich, S., 213(30), 216(30), 248 Danforth, W. H., 232(32), 240(31), 243 (32), 248 Davie, E. W., 243(109), 250 Davies, J., 42(21), 73 Davison, J., 44(103), 70(103), 75 Deal, W. C , Jr., 210 DeCastro, I. N., 103(106), 105(106), 116 DeCrombrugghe, B., 43(68), 66(22, 25), 67(22), 73, 74 DeDeken, R., 6(10), 37 DeHaan, E. J., 111(129), 117 DeLange, R. J., 232(33), 233(33), 234 (33), 243(33), 248 DeMayer, L., 173(35), 176(35), 187(35), 207 Demerec, M., 21, 37 DeMoss, R. D., 102(97), 104(97), 116 Denton, M. D., 170(103), 209 Desjardins, P. R., 139(30, 31), 162 Dessen, P., 81(18), 87(53), 93(53), 114,115 Deuticke, B., 229(49), 248 Devijlder, J. J. M., 185(26, 27), 186(28), 192(27), 207 DeVincenzi, D. L., 214(34), 248 DeWulf, H., 242(70), 249 diFranco, A., 95(68), 115 Dus, R., 158(25, 142), 162, 165 Dimroth, P., 131(32), 162 diPrisco, G., 99(81), 115 Dobrogosz, W. J., 53(23), 73 Dorsey, J. A., 136(33), 157(72), 158(117), 160(33), 162, 164 Dreisbach , R . H. , 229(49) , 248 Drochmans, P., 238(166), 251 Duba, C , 151(135), 155(135), 165 Dubois, E., 24(17a), 32 (lla),S7, 38 Duewer, T., 213(17), 214(17), 222(17), 247

256

AUTHOR INDEX

Dunn, J. J., 43(14), 45(14), 73 Durchschlag, H., 192(29, 30), 207 E Eakin, R. E., 121(34), 162 Ebashi, S., 233(122), 239(36, 37, 122), 247 (35, 36), 248, 250 Echols, H., 44(103), 70(103), 75 Eckfeldt, J., 194(31), 195(31), 196(31), 199 (31), 207 Edelstein, S. J., 177(32), 178(32), 180(32), 182(32), 184(32), 207 Edwards, J. B., 129(145), 134(145), 165 Eggerer, H., 154(87), 156(87), 164 Eigen, M., 168(33), 173(34, 35, 36), 176 (35), 181(33, 34), 185(73), 187(35, 73), 188(33), 197(33, 73), 203(33), 207, 208 Eisele, B., 192, 207 Eisen, H., 70(24), 71(24), 73 Eisenberg, H., 79(3, 13, 14), 81(3, 13, 14, 16), 114 Eisenhardt, R. H., 173(13), 207 Eliot, R. S., 179(96a), 209 Elovson, J., 144(125), 145(36, 125), 146 (35, 124), 162, 163, 165 Emmer, M., 43(68), 66(22, 25), 67(22), 73, 74 Endo, M., 239(36), 247(35, 36), 248 Engel, P . C , 87, 88, 115 Englard, S., ] 85(79), 209 Englesberg, E., 24(36), 38, 72(26), 73 Englund, P . T., 174(38), 208 Ennis, H. L., 60, 73 Eron, L., 63(89), 66(27), 68(27), 73, 74 F Fagan, V. M., 123(140), 165 Fahien, L. A., 84(34), 87(34), 88(55), 97 (71), 114, 115 Feldman, K , 219(38), 248 Ferdinand, W., 171(39), 208 Fincham, J. R. S., 105(111, 112), 107(111, 112), 116 Firsov, L., 219(8), 247 Fischer, E . H., 8(12), 37, 204(40), 208, 212(45, 46, 90), 213(17, 54, 89, 143, 144, 147), 214(17, 143, 144), 216(39, 40, 43), 217(47, 61, 86, 91, 146), 218 (7, 47, 62, 63, 147, 148, 149, 151), 219 (89, 127, 149), 221(76, 121, 146, 168), 222(17, 41, 46, 121, 146), 223(1), 224

(48, 60, 113), 226(42, 76), 228(42, 76, 92, 94), 233(44, 64, 95, 114), 234(64, 162a), 236(54, 121), 237(113), 238(64), 239(64), 240(60), 242(48), 243(89), 247, 248, 249, 250, 251 Fisher, H. F., 79(15), 82(19, 20, 21, 22, 23), 83, 85(41), 86(15), 87(52), 89(59), 93(52), 98(75), 114, 11δ Fisher, J. R., 171(104), 209 FJatt, J. P., 120(37), 163 Forrey, A. W., 216(40), 217(47), 218(47), 248 Fosset, M., 224(48), 242(48), 248 Francavilla, A., 111(128, 130, 131), 117 Freedman, R. B., 99(87), 116 Freicherr, V., 212(98), 250 Frieden, C , 77(1), 78(1), 79, 83(25, 26, 39, 30, 31, 32), 86(45), 87(47), 88(54), 90 (60, 61), 91, 92(61, 63), 93(63, 64, 65), 94, 95(25), 96(69), 97(29, 30, 70), 98 (66), 99(65, 85, 86, 89), 100(85, 86, 89), 101(1, 89), 111(64), 113, 114, H5, 116, 168(41), 171(41), 204(47), 206(41), 208 Furfine, C. S., 184(112), 185(79), 186(112), 209, 210

G Gallego, E., 174(74), 179(74), 185(74), 187 (74), 188(74), 190(74), 192(74), 198(74), 208 Ganguly, J., 159(38), 163 Garen, A., 41(100), 75 Garland, P. B., 135(156), 160(156, 157), 166 Geraci, G., 180(89), 181(89), 183(89), 209 Gerbach, E., 229(49), 248 Gerhart, J. C., 18(13), 37, 193(42), 194 (42), 195(15), 196(42), 197(17, 42), 199, 207, 208, 226(50), 248 Gerwin, B. I., 132(39, 64), 163 Ghosh, S., 44(103), 70(103), 75 Gibson, D . M., 120, 123(123), 139, 159 (10, 40, 41), 162, 163, 165, 166 Gibson, Q. H., 173(13, 44), 176(45), 177 (32), 178(32, 43, 45), 179, 180(32, 45, 89), 181(43, 89), 182(32, 45), 183(81a, 89), 184(32), 186(45), 205, 207, 208, 209 Gierer, A., 59, 73 Gilbert, W., 33(14), 37, 41(59), 42(29, 30), 43(58, 59), 44(58), 45(30), 47(29), 48, 49(29), 60, 73, 74

257

AUTHOR INDEX Ginsburg, A., 170(46, 103), 208, 209 Glaser, L., 222(111), 224(112), 250 Goff, C. G., 243(51), 248 Gold, A. M., 214(52), 216(52), 223(53), 2/+8 Goldberg, M . E., 214(156), 251 Goldfine, H., 102(98), 104(98), 116, 143(5, 43, 44), 162, 163 Goldin, B . R., 86(45), 99(85, 86, 89), 100(85, 86, 89), 101(89), 115, 116, 204 (47), 208 Goldman, J. K , 157(180), 160(172), 166 Gòlker, C. F., 214(129a), 250 Goldthwait, D . A., 43(2), 72 Gonzalez, C., 234(162a), 251 Goodall, D., 174(74), 179(74), 185(74), 187 (74), 188(74), 190(74), 192(74), 198(74), 208 Goto, T., 123(45), 163 Gottesman, M., 43(68), 66(22), 67(22), 73,

n

Graffe, M., 43(92), 75 Gratzer, W. B., 192, 208, 214(14), 247 Graves, D . J., 22(41), 213(54), 217(80a), 223(155), 224(55,164), 226(92), 236(54), 248, 249, 251 Green, A. A., 8(9), 37, 222(23), 247 Green, M . H., 70(32), 73 Green, N . M., 121, 163 Greengard, P., 246(56), 248 Greenspan, M . D., 147(51), 148(51), 149 (52, 53), 163 Gregolin, C., 123(47, 48, 49, 50), 124(47, 49, 50), 125(47), 126(49, 50), 133(47, 48,131), 134(50,131), 135(131), 163,165 Grenson, M., 3(3), 6(1, 2), 21(3, 40), 24(3, 17a, 17b, 40), 33(40a), 34(17b), 37, 38 Greville, G. D., 79(10), 114 Griffin, C . G . , 171(48), 208 Grillo, M. A., 18(15), 37 Gross, C., 200(110), 209 Grossman, I., 238(16), 247 Grundberg-Manago, M., 43(92), 75 Guchait, R. B., 131(32), 162 Guchhait, R. B., 131(32), 159(23), 161 (23), 162 Guidotti, G., 168(49), 177(49), 181(49), 188(49), 208 Guillory, R. J., 223(117), 250 Guirard, B . M., 216(57), 248 Günther, S., 151(135), 155(135), 165 Gurin, S., 123(18), 162

Gussin, G. N., 41(16), 73 Gutfreund, H., 180(69), 182(69), 204(50), 208

H Haavik, A. G., 141(22), 158(22), 162 Halford, S., 204(50), 208 Halleux, P., 3(16), 37 Halpern, Y. S., 103(101), 104(101), 116 Hammes, G. G., 56, 72, 173(36, 51), 176 (51, 54), 194(31, 53), 195(31, 52, 53, 55), 196(31), 197(52, 55), 198(55), 199(31, 55), 205(54), 207, 208 Hancock, W. S., 145(54), 149(52), 163 Hansen, J., 103(102), 104(102), 116 Hansford, R. G., 233(58), 248 Hanson, R. W., 120(11), 162 Hardman, J. G., 234(12), 246(12), 247 Hardman, J. K., 102(99), 116 Harmsen, B . J. M., 185(26), 207 Harrington, W. F., 184(56), 208 Harris, J. I., 184(57), 208 Hartmann, P . E., 21(11), 37 Hartridge, H., 177(58), 179(58), 208 Haschke, R. H., 204(40), 208, 224(60,113), 233(64), 234(64), 237(113), 238(64), 239(64), 240(59, 60), 248, 249, 250 Hasselberger, F . X., 224(100), 250 Hatfield, D., 72(31), 73 Hayaishî, O., 243(72), 249 Hayward, W. S., 70(32), 73 Heck, H . d'A., 168(59), 200(60, 61), 201 (60), 208 Hedrick, J. L., 213(147), 214(34), 216 (40), 217(61), 218(62, 63, 147, 148, 149), 219(149), 248, 251 Heilmeyer, L., Jr., 204(40), 208, 224(60, 113), 233(64), 234(64), 237(113), 238 (64), 239(64), 240(60), 248, 249, 250 Heinstein, P . F., 133, 135(55), 163 Hellerman, L., 86(44), 101(91), 115, 116 Helmreich, E., 212(67), 213(82), 214 (129a), 216(82), 218(65), 219(38), 222 (111), 223(81, 83), 224(68, 82, 83, 112), 230(69), 240(31), 244(66), 248, 249, 250 Henderson, T . O., 161(56, 57), 163 Herbst, M., 79(12), 81(12), 114, 150(118), 165 Hermann, R. L., 17(17), 38 Hers, H . G., 242(70), 249 Herzenberg, L. A., 61(33), 73

258

AUTHOR INDEX

Hickenbottom, J. P., 235(150), 242(150), 251 Hicks, S. E., 159(40), 163 Hiernaux, D., 24(17a), 32(lla), 37, 38 Hilger, F., 24(17b), 34(17b), 38 Hill, R. L., 143(166), 144(166, 167), 166 Hilvers, A. G., 185(27), 192(27), 207 Himes, R. H., 121(58), 163 Hirschbein, L., 247 Ho, E. S., 234(162b), 243(162b), 251 Huberman, J. A., 174(38), 208 Hochreiter, M. C , 88, 89(57, 58), 115 Hofnung, M., 72(31), 73, 241(141), 242 (141), 251 Hohorst, H. J., 232(71), 249 Holbrook, J. J., 99(84), 116 Hollenberg, C. P., 103(107), 105(107), 107 (107), 116 Holzer, H., 8(23, 24), 18(18), 38, 103 (104, 105), 105(104, 105), 116, 243(170), 251 Honjo, K., 120(148), 166 Horjo, T., 243(72), 249 Hooper, A. B., 103(102), 104(102), 116 Hopkins, N., 43(17), 70(17, 76), 73, 74 Hopper-Kessel, I., 154(87), 156(87), 164 Horiuchi, T., 41(34), 67(64), 73, 74 Home, R. W., 79(10), II4 Hosoi, K , 233(122), 239(122), 250 Hsu, R. Y., 141(59), 156(59), 157(59,184), 163, 166 Huang, C , 83, 92(63), 93(63), 95(25), 114, 115 Hubbard, D. D., 139,159(10, 41), 162,163 Hughes, R. C., 216(40), 248, 249 Hullar, T. L., 219(74), 249 Hunkeler, F. L., 233(9, 93), 234(9), 235 (150), 238(9), 239(93), 240(9), 242(150), 247, 249, 251 Hurd, S. S., 221(76), 226(42, 76), 228(42, 76), 236(75), 248, 249 Hurlbut, E. C., 230(138), 251 Huston, R. B., 233(77, 93), 239(93), 249 Huttunen, J. K., 243(78), 249 Huxley, H. E., 247(79), 249

I Ihler, G., 63(89), 74 Iida, Y., 67(64), 74 Ilgenfritz, G., 171(99), 178(99), 180(62, 99), 181 (99, 100), 182(62, 99), 208, 209

Illingworth, B., 216(80), 219(80), 223 (117), 249, 250 Ilton, M., 159(60), 163 Inoue, H., 120(148), 166 Ippen, K., 63(89), 65(35, 52), 67(35), 73, 74 Iwatsubo, M., 92(62), 95(68), 115, 202 (66), 208 Izui, K , 136, 163 J Jackson, 8., 106(116,119), 108(116, 119), 109(116), 116 Jackson, W. J. H., 168(86), 209 Jacob, E. J., 157(27, 62), 162, 163 Jacob, F., 3(19), 18(30), 21(20), 33(19), 38, 39(36), 41(37, 65, 102), 42(21), 47(53), 61(Ì02), 67, 69(36), 70(24), 71(24), 73, 74, 75, 223(118), 250 Jacob, M. L, 123(123), 165 Jacobs, R., 139(63, 95), 163, 164 Jacobson, B. E., 132(3, 39, 64), 162, 163 Jacobson, G. R., 223(155), 251 Jaenicke, R., 184(64), 192, 208 Jallon, J. M., 95(68), 115 Janin, J., 200(67, 109, 110), 201(18, 65, 67, 109), 202(66), 203(67), 207, 208, 209 Jansz, H. S., 216(80), 219(80), 249 Jeckel, R., 99(84), 116 Jevans, A. W., 159(60), 163 Jirgensons, B., 83, 114 Jobe, A., 44(10), 47(10), 48(10), 49(38), 62(10, 38), 72, 73 Johnson, J. F., 217(80a), 249 Johnson, P., 83(33), II4 Jones, O. W., 43(39), 47(39), 73 Jornvall, H., 79(8), II4 Joshi, V. C., 157(65), 158(65, 120), 160 (119), 163, 165 Jutting, G., 121(88), 129(88), 164 Jovin, T. M., 174(38), 205(68), 208 K Kahn, V., 224(80b), 249 Kamen, R. I., 70(94, 95), 71(95), 75 Kameyama, T., 67(64), 74

259

AUTHOE INDEX

Kaplan, N. 0., 84(35), 85(39), 87(49), 88, 97(49), 114, Uà Karpatkin, S., 223(81), 249 Karr, G. M., 184(56), 208 Kastenschmidt, J., 213(82), 216(82), 223 (83), 224(82, 83), 249 Kastenschmidt, L. L., 213(82), 216(82), 223(83), 224(82, 83), 249 Kato, I., 243(72), 249 Katsuki, H., 136(61), 163 Katz, J., 120(129), 165 Keech, D. B., 122(136), 165 Keller, P. J., 213(84), 213(85), 237(84), 249 Kellet, G. L., 180(69), 182(69), 208 Kemp, R. G., 232(33), 233(33), 234(33), 243(33), 248 Kennan, A. L., 159(29), 161(21, 29), 162 Kent, A. B., 216(40, 43), 217(86), 226(94), 248, 249 Kepes, A., 53(15), 60(40), 73 Khan, R. P., 120(115), 165 Kihara, H. K , 43(44), 73 Kilburn, E., 137, 138, 139(63, 96), 163,164 Kim, S. J., 81(17), 114 King, C. A., 235(132), 250 King, K. S., 83(30), 97(30), 114 Kingdon, H. S., 170(103), 209, 243(87), 249 Kirk, P. R., 110(125), 117 Kirschner, K., 168(71), 169, 174(74), 179 (74), 185(73, 74), 187(73, 74), 188(74), 190(74), 191(70, 72), 192(29, 30, 74), 197 (73), 198(74), 205(72), 207, 208 Klein, H. P., 120, 163 Kleinschmidt, A. K., 123(47, 49, 50, 67), 124(47, 49, 50, 67), 125(47), 126(49, 50, 67), 133(47), 134(50), 163 Klemm, A., 44(6), 62(6), 72 Kleppe, K., 213(30), 216(30), 248 Klossen, G. R., 106(119), 108(119), 116 Knappe, J., 121(68, 69, 88), 122(69), 129 (68, 88), 163, 164 Knivett, V. A., 148(70), 163 Knof, S., 184(64), 208 Koch, A., 42(41), 45(42), 73 Koh, P., 226(42), 228(42), 248 Kornacker, M. S., 120(71), 164 Kornberg, A., 174(38), 208 Koshland, D. E., Jr., 47, 73,109(122), 116, 167(75, 76), 168(75, 76, 77), 169(25, 75, 77), 170(80), 171(75, 76), 178(75, 77),

185(23, 24), 186(23), 188(75, 77), 190, 197(75, 77), 201(75, 77), 203(75, 77), 206(75, 77), 207, 208, 209 Kratky, O., 150(118), i05,192(29,30),207, 214(129a), 250 Krebs, E. G., 8(12), 37, 212(45, 90), 213 (54, 89), 216(40, 43), 217(86, 91), 219 (89), 221(121, 168), 222(41, 121, 171), 226(92, 94), 232(33, 88), 233(9, 33, 44, 77, 93, 95, 114), 234(9, 33, 129, 162a, 162b, 163), 235(88, 132,132a, 133, 150), 236 (54, 121), 238(9), 239(93), 240(9), 242(150), 243(20, 33, 89, 162b), 247, 248, 249, 250, 251 Krebs, H. A., 112(133, 134), 117 Krimsky, I., 184(92), 209 Kroeplin-Rueff, L., 156(134), 165 Kumar, S., 157(72), 164 Kuman, A., 249 Kuno, H., 43(44), 73 Kustin, K., 173(78), 176(78), 209 L Lachance, J. P., 121(88), 129(88), 164 Landon, M., 79(5), 80(5), 99(83), 100(5), 114, 116 Lane, M. D., 122(73), 123(47, 48, 49, 50, 67), 124(47, 49, 50, 67), 125(47), 126 (49, 50, 67), 129(145), 131(32), 133(47, 48, 74, 131), 134(48, 50, 145), 135(131), 159(26), 162, 163, 164, 165 Langdon, R. G., 123(75), 164 Langley, T. J., 79(5), 80(5), 100(5), II4 Lardy, H. A., 120(186), 166 Larner, J., 243(161), 251 Larrabee, A. R., 141(163), 142(163), 143 (163), 145(162), 146(158), 152(76), 161 (158), 164, 166 Lata, M., 105(113, 114,115), 107(113, 114, llò),116 Lazdunski, C , 204(77a), 209 Lazdunski, M., 204(77a), 209 LeBras, G., 201(108), 209 Leder, P., 43(63), 74 Leech, R. M., 110(125), 117 LéJohn, H. B., 105(108, 109, 110), 106 (116, 117, 118, 119, 120, 121), 107(108, 109, 110), 108(116, 117, 118, 119, 120, 121), 109(116, 118, 120, 121), 110(121), 116 Lelange, R. J., 235(133), 250

260

AUTHOR INDEX

Leloir, L. F., 223(97), 249 Lelouchier-Dagnelie, H., 14(6), 37 Lennarz, W. J., 148(77), 164 Lerch, I., 156(134), 165 Leveille, G. A., 120(78), 164 Levitzki, A., 109(122), 116, 170(80), 209 Levy, H. R., 133(105), 164 Liberati, M., 146(158), 161(158), 166 Liebig, J., 212(98), 250 Lies, K , 8(24), 38 Light, R. J., 148(77), 164 Lin, S. Y., 47(45), 73 Lipmann, F., 235(154), 239(37), 248, 251 Lipscomb, W. N., 172(117), 210 Listowsky, L, 185(79), 209 Lochmuller, H., 122(183), 166 Lohmann, K., 229(99), 250 Lonberg-Holm, K. K., 173(13), 207 Long, C. W., 170(80), 209 Long, R. W., 160(122), 165 Loomis, W. F., 66(46), 73 Lorch, E., 121(88), 129(88), 164 Lou, H. F., 17(17), 38 Love, D . S., 233(95), 249 Lowenstein, J. M., 120(71), 164 Lowry, H. O., 224(100), 250 Lubin, M., 60, 73 Lumry, R., 180(84), 181(84), 209 Lund, P., 112(133), 116 Lust, G., 136(79), 160(79), 164 Lynen, F., 119(85), 121(58, 68, 85, 88), 122 (73, 103, 111, 183), 123(102, 103, 109), 129(68, 88), 133(103, 109, 112), 135(16, 17, 109), 136(79, 109), 141(88, 85), 144 (179), 150(80, 85, 114, 118), 151(80, 83, 84, 85, 86, 135, 147), 152(80, 85, 176, 179), 153(85), 154(81, 83, 85, 87, 89), 155(85, 89, 135), 156(80, 87, 134, 146), 159(110), 160(79), 162, 163, 164, 165, 166, 237(101), 250 Lyon, J. B., 223(102), 232(32, 103), 243 (32), 248, 250

M McCarty, E. D., 159(60), 163 McClintock, D . K., 194(82), 209, 233 (104), 250 McConnell, H. M., 168(81), 183(81, 87), 209 McCrea, B . E., 105(110), 107(110), 116

McDaniel, E . G., 152(76), 164 McGregor, L. L., 82(19, 20), 85(41), 86, 98(75), 114, H5 Machattie, L., 63(89), 74 McNeill, J. J., 161(56, 57), 163 MacQuarrie, R., 183(81a), 209 Madsen, N . B., 213(106), 215(105), 216 (4, 5, 6), 223(105), 224(105, 108), 229 (105), 247, 250 Magasanik, B., 2(32), 38, 65(48), 66(46, 90), 73, 74 Mahler, R., 223(140), 251 Majerus, P. W., 119(97), 135(97), 137,138, 139(63, 95, 96), 141(97, 163), 142(92, 97, 163), 143(97, 163), 144(90, 91, 93, 94), 146(6), 147(7), 150(97), 154(7, 97), 162, 163, 164, 166 Makman, R. S., 66(49), 73 Malcolm, A. D . B., 101(96), 116 Mandelstam, J., 2(21), 38 Maragoudakis, M . E., 140(98, 99, 100), 164 Margolis, S. A., 159(23), 161(23), 162 Markus, G., 194(82), 209, 233(104), 250 Marier, E., 79(2), II4 Marshall, G. R., 145(54), 163 Martello, O. J., 243(109), 250 Martin, D . B., 123(101, 160, 161), 126 (160, 161), 133(101, 161), 164, 166 Martin, R. G., 18(22), 38, 69, 73 Matsuhashi, M., 122(103), 123(102, 103), 133(103), 159(110), 164, 165 Matsuhashi, S., 122(103), 123(102, 103), 133(103), 164 Matsumura, S., 141(19), 162 Matthes, K. J., 123(1), 158(2), 162 Mayer, S. E., 243(78), 249 Mayor, F., 103(106), 105(106), 116 Mecke, D., 8(23, 24), 38, 243(170), 251 Meighen, E . A., 184(83), 209 Melamed, M . D., 79(5), 80(5), 100(5), 114 Merlevede, W., 241(110), 250 Meronk, F., 72(26), 73 Merrifield, R. B., 145(104), 164 Messenguy, F., 6(25, 26, 27, 28), 7(25), 8 (25), 9(25), 10(25, 28), 11(25, 28), 12 (28), 13(25, 28), 14(25, 26), 15(25, 28a, 32a), 16(28, 28a), 17(28a), 18(28a), 19 (28a), 20 (28a), 35(28, 28a), 38 Metzger, B . E., 222(111), 224(112), 250

261

AUTHOR INDEX

Meuser, R., 106(121), 108(121), 109(121), 110(121), 116 Meyer, F., 224(60, 113), 233(64), 234(64), 237(113), 238(64), 239(64), 240(60), 248, 249, 250 Meyer, W. L., 233(95, 114), 249, 250 Meyerhof, 0., 212(115), 250 Middelhoven, W. J., 32(29), 88 Miller, A. L., 133(105), 164 Miller, J. H., 40(51), 63(70, 72), 65(35, 52), 66(90), 67(35, 77), 78, 74 Milroy, T. H., 212(116), 250 Mizukami, H., 179(96a), 180(84), 181(84), 209 Mohr, S. C , 194(31), 195(31), 196(31), 199(31), 207 Mommaerts, W. F. H. M., 223(117), 250 Monod, J., 3(19), 3(31), 18(30), 33(19), 88, 39(36), 41(37, 65, 102), 47(53, 55), 53 (15, 54, 97), 60(11), 61(55, 102), 66(97), 67, 69(36), 72, 73, 74, 75, 95, 115, 167 (85), 168(85), 169(85), 171(85), 182(85), 185(85), 188(85), 191(85), 192, 194(85), 197, 198(85), 201(85), 202, 205(85), 206 (85), 209, 214(156), 215(157), 223(118), 224(157), 250, 251 Morgan, H. E., 215(123), 227(119), 229 (119, 123), 230(119), 250 Morikawa, M., 136(61), 163 Morris, H. P., 139(95), 164 Moses, V., 53(73), 74 Moss, J., 123(67), 124(67), 126(67), 163 Müller, W., 51, 74 Müller-Hill, B., 33(14), 87, 40(51), 41(57, 59), 42(29, 30), 43(58, 59), 44(6, 58), 45 (30), 47(29), 48(6, 29), 49(29, 60), 53 (60), 60, 62(6), 63(70), 72, 78, 74 Muesing, R. A., 157(72), 164 Muir, L. W., 224(48), 242(48), 248 Munday, J. S., 97(72), 115 Myers, G. L., 48, 49(62), 74

N Nakanishi, S., 123(106), 124(106), 126 (106), 138, 164 Narahara, H. T., 250 Neet, K. E., 167(77), 168(77), 169(77), 178(77), 188(77), 197(77), 201(77), 203 (77), 206(77), 208 Neidhardt, F. C , 2(32), 88

Nervi, A. M., 129(8), 131(107), 132(107), 135(8), 162, 164 Nester, E. W., 20(8), 87 Newby, R. F., 42(81, 82), 43(81), 49(82), 50(82), 51(81), 53(82), 57(82), 66(82), 74 Newsholme, E. A., 232(135, 158), 251 Nichol, L. W., 168(86), 209 Nielsen, L. D., 224(48), 242(48), 248 Nirenberg, M., 43(63), 74 Nishida, M., 101(92, 95), 116 Nishizuma, Y., 243(72), 249 Nixon, J. E., 139(108), 157(108, 116), 158 (108, 116, 117), 164, 165 Nolan, C , 221(121), 222(121), 236(121), 250 Notani, G., 44(20), 73 Novick, A., 41(86), 74 Novoa, W. B., 221(121), 222(121), 236 (121), 250 Nulty, W. L., 145(54), 163 Numa, S., 122(111), 123(45, 102, 106, 109, 113), 124(106), 126(106), 133(109, 112), 135(109), 138, 159(110), 168, 164, 165

o O'Brien, J. R. P., 179(25a), 207 Oesterhelt, D., 150(114, 118), 154(89), 155 (89), 156(146), 164, 165 Özand, P., 250 Ogawa, S., 183(87, 88), 209 Ohga, Y., 235(153), 251 Ohshima, Y., 41(34), 67, 73, 74 Ohtsuki, I., 239(36), 247(36), 248 Oliveira, R. J., 224(55), 248 Oison, E. B., 159(23), 161(23), 162 Olson, J. A., 85(40), 87(40), 101(40), 115 Oison, J. S., 177(32), 178(32), 180(32), 182 (32), 184(32), 207, 209 Orgel, L. E., 41(9), 72 Osber,M.P.,223(53),£4