Lysosomal Pathways of Protein Degradation (Molecular Biology Intelligence Unit)

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MOLECULAR BIOLOGY INTELLIGENCE UNIT 11

J. Fred Dice DICE MBIU 11

Lysosomal Pathways of Protein Degradation


MOLECULAR BIOLOGY INTELLIGENCE UNIT 11

Lysosomal Pathways of Protein Degradation J. Fred Dice, Ph.D. Department of Physiology Tufts University School of Medicine Boston, Massachusetts, U.S.A.

LANDES BIOSCIENCE GEORGETOWN, TEXAS U.S.A.

EUREKAH.COM AUSTIN, TEXAS U.S.A.

LYSOSOMAL PATHWAYS OF PROTEIN DEGRADATION Molecular Biology Intelligence Unit EUREKAH.COM LANDES BIOSCIENCE

Copyright ©2000 EUREKAH.COM All rights reserved. No part of this book may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Printed in the U.S.A. Please address all inquiries to the Publishers: EUREKAH.COM/ Landes Bioscience, 810 South Church Street Georgetown, Texas, U.S.A. 78626 Phone: 512/ 863 7762; FAX: 512/ 863 0081

ISBN: 1-58706-003-5

Library of Congress Cataloging-in-Publication Data Dice, J. Fred, 1947Lysosomal pathways pf protein degradation/J.Fred Dice. p. cm. -- (Molecular biology intelligence unit) ISBN 1-58706-003-5 (alk. paper) 1. Lysosomes. 2. Proteins 3. Endocytosis. I. Title. II Series. [DNLM: 1. Endocytosis--physiology 2. Lysosomes--physiology. 3. Proteins--metabolism. QH603.L9 D546L 2000] QH603.L9 D546L 2000 571.6'55--dc21 99-027040 CIP

CONTENTS 1. Introduction ........................................................................................... 1 Methods Used to Study Lysosomes ....................................................... 1 Terminology ......................................................................................... 2 Acidification of Lysosomes .................................................................... 3 Biogenesis of Mammalian Lysosomes .................................................... 7 Biogenesis of the Yeast Vacuole ........................................................... 11 Degradation of Lysosomal Proteins ..................................................... 12 Future Directions of Research ............................................................. 13 2

Degradation of Endocytosed and Plasma Membrane Proteins ............. 18 Overview ............................................................................................. 18 Methods Used to Study Endocytosis ................................................... 21 Mechanisms of Endocytosis in Mammalian Cells ................................ 23 Degradation of Plasma Membrane Receptors ...................................... 29 Receptor-Mediated Endocytosis in Yeast ............................................. 32 Future Directions of Research ............................................................. 33

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Lysosomal Degradation of Proteins in the Secretory Pathway: Crinophagy ........................................................................... 39 Overview ............................................................................................. 39 Methods Used to Study Protein Secretion and Crinophagy ................. 39 Constitutive Secretion Pathways .......................................................... 43 Regulated Secretion Pathways .............................................................. 45 Neurotransmission and Synaptic Vesicle Dynamics ............................. 48 Crinophagy ......................................................................................... 51 Future Directions of Research ............................................................. 53

4. Degradation of Intracellular Proteins by Macroautophagy ................... 57 Overview ............................................................................................. 57 Methods Used to Study Macroautophagy ............................................ 59 Tissue Specificity of Macroautophagy .................................................. 61 Mechanisms of Macroautophagy ......................................................... 62 Regulation of Macroautophagy ........................................................... 63 Substrate Selectivity ............................................................................. 69 Future Directions of Research ............................................................. 70 5. Degradation of Intracellular Protein by Microautophagy ..................... 75 Overview ............................................................................................. 75 Methods Used to Study Microautophagy ............................................ 75 Mechanisms of Microautophagy .......................................................... 78 Regulation of Microautophagy ............................................................ 82 Future Directions of Research ............................................................. 82

6. Selective Pathway for Degradation of Cystolic Proteins by Lysosomes ....................................................................................... 85 Overview ............................................................................................. 85 Background ......................................................................................... 85 Methods Used for the Study of Selective Lysosomal Proteolysis ........... 86 RNase A as a Substrate for Selective Lysosomal Proteolysis .................. 90 The KFERQ Motif ............................................................................. 90 Reproduction of the Selective Pathway of Lysosomal Proteolysis In Vitro ............................................................................................ 92 The Receptor in the Lysosomal Membrane ......................................... 93 Possible Roles of Other Lysosomal Membrane and Matrix Proteins ......................................................................... 98 Regulation of the Selective Lysosomal Proteolytic Pathway ................. 99 A Working Model for the Selective Lysosomal Degradation Pathway .................................................................... 100 Future Directions of Research ........................................................... 101 7. Concluding Remarks .......................................................................... 105 Another Pathway for the Delivery of Cytosolic Proteins to the Yeast Vacuole ...................................................................... 105 Index .................................................................................................. 107

PREFACE

P

roteins are continually degraded and resynthesized within cells, and this constant turnover allows the cell to adjust concentrations of different proteins under different physiological conditions. Protein turnover also prevents proteins from accumulating a variety of posttranslational modifications that could be deleterious to protein function. Protein degradation can also serve as a quality control system that removes proteins which have not folded properly or have not assembled into appropriate macromolecular complexes. Finally, protein degradation provides a source of amino acids to be used for energy or for the synthesis of essential proteins under conditions where amino acids from the diet are in short supply such as during fasting. I have studied intracellular protein degradation since I began graduate school at Stanford University in 1969. My thesis advisor, Robert T. Schimke, M.D., made several important contributions to this area of research including the demonstration that the degradation rates of different proteins within a single organelle could be dramatically different. This finding indicated that organelles were generally not degraded as a unit. Bob also showed that crucial regulatory proteins were short-lived and that their rapid degradation rate allowed their intracellular levels to be adjusted quickly. The popularity of research on intracellular protein degradation has increased dramatically over the past 30 years, and it is now clear that multiple pathways of proteolysis exist within cells. Lysosomes are acidified hydrolytic organelles surrounded by a unit membrane. The lysosomal hydrolases include proteases and peptidases, nucleases, phosphatases, lipases, and glycosidases. Lysosomes are dynamic organelles which can fuse with each other and with certain other organelles and show dramatically altered morphology under particular physiological and pathological conditions. Lysosomes are able to take up and degrade intracellular proteins by a variety of different mechanisms that are reviewed in this book. The importance of lysosomes in the degradation of intracellular proteins varies depending on the cell type. Lysosomes are responsible for most of the degradation of long-lived proteins in liver and in many other tissues, but lysosomes play a more limited role in the degradation of proteins in skeletal muscle and lymphocytes. The yeast vacuole is homologous to the mammalian lysosome in many ways, and vacuolar proteases are responsible for much of the protein degradation under conditions of nitrogen or carbon starvation and during sporulation. The major nonlysosomal proteolytic pathway in both yeast and mammalian cells is the ubiquitin/proteasome pathway. This important pathway of proteolysis is responsible for the degradation of many short-lived normal and abnormal proteins and is also responsible for the degradation of certain long-lived proteins.

This highly regulated pathway of proteolysis is also required for the maturation of autophagic vacuoles. It is not yet known whether ubiquitination of critical lysosomal components is required for their activation or whether substrate proteins require ubiquitination for their uptake by and degradation within lysosomes. Uncovering the mechanisms and the physiological importance of the linkage between the ubiquitin/ proteasome and lysosomal degradation pathways remains a critical area for future research.

ACKNOWLEDGMENTS I thank my colleague, Ana Maria Cuervo, for critical reading of an early draft of this book and for many valuable suggestions for improvement. I also thank the many talented individuals who have contributed to the research in my laboratory concerning the selective pathway of lysosomal proteolysis (Chapter 6). These individuals include: Carlos Walker, Meg McCrystal, Alex Cardiel, Betsy Byrne, Emma Hess, Fred Kalish, Nora Chovick, Felipe Samaniego, Faye Berry, Joe Auteri, Annabelle Okada, Victor Bochaki, Betty Bourret, Nicki Neff, Paul Miao, Peter Netland, Mary Ann McElligott, Jon Backer, Liz Spencer, HuiLing Chiang, Eric Gulve, Joe Berger, Mia Meyer, Steve Goff, Sharla Russell, Laura Terlecky, Stan Terlecky, Sabine Freundlieb, Tracy Olson, Lois Isenman, Charles Plant, Melissa KirvenBrooks, Simon Wing, Fernando Agarraberes, Marilyn Negron, Sandra Hayes, Elizabeth Frutiger, Melissa McMahon, Amy Majesty and Ana Maria Cuervo. I have also had the good fortune to have collaborations with Erwin Knecht, Wei Hu, Bing Lim, Heinz Hildebrand, Ernst Bomhard, and Junor Barnes. Visiting scientists contributed to the research and training in my laboratory. These have included Roger Dean, Letitia Beard, and the late Hal Segal. I also thank Edward F. C. Blommaart for sending me a copy of his Ph.D. thesis, Regulation of Hepatic Autophagy by Amino Acid Dependent Signal Transduction, reporting research results he carried out in the laboratory of A. J. Meijer in the Department of Biochemistry, Academic Medical Center, University of Amsterdam, The Netherlands. It arrived in the mail on the day I began collecting articles and reading about macroautophagy, and it was extremely helpful. The research from my laboratory has been supported by the National Institutes of Health by grants from the National Institute of Arthritis and Metabolism (AM21104) and the National Institute on Aging (AG02783, AG07472, and AG06116).

DEDICATION This book is dedicated to Robert T. Schimke, M.D., my Ph.D. thesis advisor at Stanford University. Bob instilled in me a love of science, both for research and for teaching. He has been an exemplary role model for dozens of scientists who have worked with him, and he continues to be a world class scientist and an exceptionally courageous human being.

ABBREVIATIONS ALP: AP-1: AP-2: AP-3: apg: API: AR: ARF: BiP: cAMP: cdM6PR: CHO: ciM6PR: CLIP-170: COPI: COPII: CPY: CURL: cvt: E1: EGF: EGFR: end: Eps15: ER: ErbB-4: ERGIC: F-ATPase: FBPase: Gi: GAPDH: GERL: Hsc73: Hsp70: ICG: IGF-II: IR:

alkaline phosphatase adaptor protein 1 complex adaptor protein 2 complex adaptor protein 3 complex autophagy mutants aminopeptidase I asialoglycoprotein receptor ADP-dependent ribosylation factor binding protein in the endoplasmic reticulum lumen; also referred to as glucose-regulated protein of 78 kilodaltons; an hsp70 family member cyclic 3′-5′ adenosine monophosphate cation-dependent mannose-6-phosphate receptor Chinese hamster ovary cation-independent mannose-6-phosphate receptor cytoskeleton-localized intermediate protein of 170 kilodaltons coatomer protein complex I coatomer protein complex II carboxypeptidase Y compartment for uncoupling receptor and ligand cytoplasm-to-vacuole targeting mutants ubiquitin activating enzyme epidermal growth factor epidermal growth factor-receptor endocytosis mutants epidermal growth factor-receptor protein substrate of 15 kilodaltons endoplasmic reticulum heregulin receptor endoplasmic reticulum-Golgi intermediate compartment F1F0-ATP synthase fructose-1,6-bisphosphatase inhibitory GTPase glyceraldehyde-3-phosphate dehydrogenase Golgi-endoplasmic reticulum-lysosome complex constitutively-expressed heat shock protein of 73 kilodaltons heat shock protein of 70 kilodaltons intercisternal granules insulin-like growth factor II insulin receptor

ISV: kDa: LAMP: LAP: LDL: LDLR: LIMP II: LGP96: ly-hsc73: M6P: M6PR: MAP: NEM: NSF: pag: PI3 kinase: POMC: PrA: PrB: rab: RER: RNase A: RNase S-peptide: RNase S-protein: SDS-PAGE: SER: SNAP: SV: TGN: TR: TSH: t-SNARE: V-ATPase: vid: vps: v-SNARE: VSV:

immature secretory vesicle kilodalton lysosome-associated membrane protein lysosomal acid phosphatase low density lipoprotein low density lipoprotein receptor lysosomal integral membrane protein II lysosomal glycoprotein of 96 kilodaltons lysosomal hsc73 mannose-6-phosphate mannose-6-phosphate receptor multiple antigen protein N-ethylmaleimide N-ethylmaleimide-sensitive factor peroxisomal autophagy mutants phosphatidylinositol-3 kinase proopiomelanocortin protease A protease B ras-related GTPase rough endoplasmic reticulum bovine pancreatic ribonuclease A amino acids 1-20 of bovine pancreatic ribonuclease A amino acids 21-124 of bovine pancreatic ribonuclease A sodium dodecylsulfate polyacrylamide gel electrophoresis smooth endoplasmic reticulum soluble NSF attachment protein secretory vesicle trans-Golgi network transferrin receptor thyroid stimulating hormone target membrane soluble N-ethylmaleimide sensitive attachment receptor vacuolar ATPase vacuolar import defective mutants vacuolar protein sorting mutants vesicle membrane soluble N-ethylmaleimide sensitive attachment receptor vesicular stomatitis virus

CHAPTER 1

Introduction

L

ysosomes are organelles that contain many hydrolases with an acidic optimum pH.1-7 Lysosomes constitute 1-15% of the total cell volume and total cell protein in most mammalian cells. The existence of such organelles was suggested more than 100 years ago by the work of Metchnikoff and others concerning phagocytosis by protozoa.1,2 They observed that bacterial aggregates could be taken up by certain protozoa and were delivered to other regions of the cell, now known to be lysosomes or vacuoles, where they were digested. Our more recent understanding of lysosomes has come from morphological studies at the level of the electron microscope as well as from biochemical fractionation studies of organelles contained in rat liver. The studies of Novikoff 3,4 and de Duve5,6 have been especially important. Novikoff showed that lysosomes could be extremely heterogeneous in morphology, but their common feature was to contain multiple acid hydrolases. de Duve showed that the particulate fraction of cell homogenates that biochemists routinely prepared to study mitochondria actually contained two other organelles as well— lysosomes and peroxisomes. His group was able to separate these organelles by sucrose density gradients (Fig. 1.1). The key features of lysosomes are that they contain many different acid hydrolases (Table 1.1) at their final site of intracellular localization,1,2,7 and that they are acidified organelles.1,2,7,8 Other organelles are also acidified (for example, endosomes9 and Golgi10), and certain hydrolases may be targeted to endosomes in some cell types.1,11,12 These considerations have led some researchers to refer to the entire Golgi-endoplasmic reticulumlysosome complex as a single entity (GERL1,13), or as endosomes/lysosomes,1,14,15 or the central vacuolar system.1,16 However, I will use the definition of lysosomes originally supplied by de Duve and Wattiaux5 and more recently refined by Kornfeld and Mellman.7 This definition emphasizes that lysosomes are a terminal degradative compartment that represents the final destination not only of lysosomal hydrolases but also of a significant fraction of all endocytosed proteins. This functional definition avoids the confusion caused by the variable morphology of lysosomes even within a single cell,1,2,7 and it also allows for the transient presence of lysosomal hydrolases in other compartments along the biosynthetic pathway of lysosomes.7

Methods Used to Study Lysosomes The analysis of lysosomes has relied on a combination of morphological, biochemical, and cell biological approaches. The morphological approaches have included analyses by electron microscopy and the use of cytochemical stains to identify structures that contain

Lysosomal Pathways of Protein Degradation, by J. Fred Dice, Ph.D. ©2000 EUREKAH.COM

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Fig. 1.1. Density gradient separation of lysosomes, mitochondria, and peroxisomes. Rat livers were homogenized and centrifuged at 12,000 x g for 10 min. The pellet was layered over a sucrose gradient ranging from 10-45% and centrifuged at 45,000 x g for 16 hr. Fractions were collected from the top of the tube, and their densities were measured (- - - - ). Enzymatic assays were also done for markers of lysosomes (acid phosphatase), mitochondria (cytochrome oxidase), and peroxisomes (urate oxidase).

acid hydrolases. Such studies have greatly increased our understanding of lysosomal structure and heterogeneity.1,3,13 Biochemical analyses with isolated lysosomes usually rely on separation from mitochondria and peroxisomes based on slight density differences (Fig. 1.1). A recent example has been the development by Madden and Storrie17 of the purification of lysosomes from cells in culture. In this case the lysosomes are separated from other organelles using two consecutive density step-gradients (Fig. 1.2). The lysosomes are forced to migrate to their equilibrium density down a density gradient in the first step and up through a density gradient in the second step. These biochemical studies with purified lysosomes have been crucial in identifying components of lysosomes and in the study of certain aspects of lysosomal function. The most convincing studies often combine morphological and biochemical approaches to establish, for example, the pathway of delivery of a lysosomal enzyme to the lysosome (see below). Cell biological studies include the use of living cells in culture and pH-sensitive dyes to measure intralysosomal pH.8,18 Also, laser scanning confocal microscopy combined with immunolocalization has proven useful in documenting that a particular protein, in fact, resides in lysosomes.7,19

Terminology The names of lysosomes and lysosome-related organelles are summarized in Table 1.2. Primary lysosomes are defined as membrane-bound, acidic organelles that contain the battery of hydrolases but not yet any substrate material to be degraded.1,2 Secondary lysosomes

Introduction

3

Table 1.1. Mammalian lysosomal enzymes and proteins Proteases and Peptidases cathepsin D, cathepsin B, cathepsin H, cathepsin L, tripeptidyl peptidase, dipeptidyl peptidase I, dipeptidyl peptidase II, arginyl aminopeptidase, carboxypeptidase A, carboxypeptidase B, prolyl carboxypeptidase, tyrosyl carboxypeptidase, dipeptidase I, dipeptidase II Nucleases ribonuclease II, deoxyribonuclease II, 5′-exonuclease, acid phosphatase, acid phosphodiesterase, pyrophosphatase, nucleoside triphosphatase Lipases triacylglycerol lipase, phospholipase A1, phospholipase A2, acylsphingosine deacylase, phosphatidate phosphatase, sphingomyelin phosphodiesterase, sphingomyelin phosphodiesterase activator proteins Glycosidases α-L-fucosidase, α-galactosidase, β-galactosidase, α-glucosidase, β-glucosidase, α-Nacetylgalactosaminidase, α-N-acetylglucosaminidase, β-glucuronidase, α-L-iduronidase, α-mannosidase, β-mannosidase, neuraminidase, β-aspartylglucosylaminidase, chondroitin-6sulfatase, heparin sulfamatase, iduronosulfatase, sulfatase A, sulfatase B, hyaluronidase, heparin endoglucuronidase, heparin sulfate endoglycosidase, lysozyme Other V-ATPase, amino acid transporters, receptors (LGP96) For V-ATPase, vacuolar ATPase (see Chapter 2); LGP96, lysosomal glycoprotein of 96 kilodaltons (see Chapters 2 and 6).

are primary lysosomes that have also received substrates.1,2 These substrates may be derived from inside or outside the cell. If the material is from outside the cell, the lysosomal digestion process has been called heterophagy.1,2 If the material is from inside the cell, the digestion is called autophagy.1,2 The same types of primary lysosomes are involved in heterophagy and autophagy. As Figure 1.3 illustrates, there are several different ways that intracellular proteins are delivered to lysosomes for degradation, and these processes are the major subject matter of this book. Secondary lysosomes that have digested many substrates but now contain largely indigestible material or digested products that cannot readily pass through the lysosomal membrane are called dense bodies, residual bodies, or telolysosomes.1 Early and late autophagosomes refer to intermediates in the process of macroautophagy. These structures are collectively called autophagic vacuoles.

Acidification of Lysosomes The pH within lysosomes can be measured with pH-sensitive dyes such as fluorescein linked to inert material. When cultured fibroblasts are allowed to endocytose fluoresceindextran, and the cells are further incubated without fluorescein-dextran for several hours so that all the dye within the cell is within lysosomes, the excitation-emission pattern indicates that the lysosome is at pH 5.2-5.6.8,20 Furthermore, incubation of the fibroblasts with a weak base such as chloroquine or ammonium chloride causes the acidified lysosomes to become neutral.20


4

Fig. 1.2. Step-gradients to purify lysosomes from cultured cells. After cell homogenization and centrifugation, the postnuclear supernatant (PNS) is layered on top of density step-gradients as indicated. Mtz = metrizamide. After centrifugation, the Golgi, light mitochondria/lysosomes (Lt Mito/Lyso), and mitochondrial (Mito) fractions accumulate at the indicated interfaces. The light mitochondria/lysosomes are then transferred to the bottom of a second density step-gradient. After centrifugation the mitochondria (Mito) and lysosomes migrate to the indicated interfaces. This technique was originally developed by Madden and Storrie using Chinese hamster ovary (CHO) cells.17

Table 1.2. Terminology used for lysosomes and lysosome-related structures Term

Definition

lysosome

A degradative compartment surrounded by a single membrane and containing hydrolases that operate optimally at acidic pH; the final site of delivery of most endocytosed proteins.

vacuole

The lysosome of yeast, fungi, plants, and protozoa.

primary lysosome

A newly made lysosome that contains hydrolases but no substrates.

secondary lysosome

A lysosome that contains both hydrolases and substrates.

residual body

A secondary lysosome that contains indigestible material or digestion products that cannot exit the organelle; also called dense body or telolysosome.

heterophagosome

A secondary lysosome that has received material to digest that originated from outside the cell.

autophagosome

A secondary lysosome that has received material to digest that originated from inside the cell.

early autophagosome

A double membrane surrounding an area of cytoplasm. The membrane contains no lysosomal membrane proteins.

late autophagosome

A double membrane surrounding an area of cytoplasm. The membrane contains V-ATPase, and the organelle is acidified.

autophagic vacuole

Refers to both early and late autophagosomes.

Introduction

5

Fig. 1.3. Lysosomes can internalize cellular proteins by many different pathways. Lysosomes can degrade endocytosed proteins (END) and secreted proteins in secretory vesicles (SV). Lysosomes can also degrade areas of cytoplasm that are first sequestered into double-membraned autophagic vacuoles (AV) by a process called macroautophagy. Lysosomes can also internalize smaller bits of cytoplasm through vesiculation of the lysosomal membrane in a process called microautophagy which may result in the formation of multivesicular bodies (MVB). Finally, lysosomes are also able to take up and degrade certain cytosolic proteins in a molecule-by-molecule fashion that is stimulated by the heat shock 73kDa molecular chaperone (HSC73-mediated). ER = endoplasmic reticulum; M = mitochondrion; L = lysosome; N = nucleus.

Lysosomal membranes are now known to contain a multi-subunit proton pump that hydrolyzes ATP and transports protons into the lysosomal lumen.21 The composition of this proton pump is very similar in mammals, yeast, and plants.21 Its overall structure resembles that of the mitochondrial F1F0-ATP synthase (F-ATPase) which synthesizes ATP using the proton gradient within mitochondria as the energy source. Just as the mitochondrial F-ATPase can be separated into membrane-associated F0 and soluble F1 multimeric halves,22 the lysosomal or vacuolar ATPase (V-ATPase) can be separated into V0 and V1 domains.21 The overall structures of F-ATPase and V-ATPase are compared in Figure 1.4. The polypeptide composition of V-ATPases appears to be A, B, and C in a 3:3:12 ratio compared to other subunits that are present as a single copy.21 There are three distinct C subunits in yeast designated C, C′, and C′′ that are each present at 4 copies per complex. The single copy subunits include proteins of 100, 40, 38, and 33 kilodaltons (kDa) of unknown function in rat liver lysosomes.23 Additional single copy proteins are found in V-ATPases localized to other organelles such as coated vesicles.21 The A subunits (73 kDa) contain the catalytic sites for ATP hydrolysis while the B subunits (58 kDa) contain noncatalytic ATP-binding sites that are presumably regulatory for activity and/or assembly of the pump.24 The C subunits (17 kDa) are responsible for proton pumping, but they are active only when assembled with the V1 complex and the single copy subunits of V0 and V1.21

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Fig. 1.4. Structural comparison of the mitochondrial F1F0-ATP synthase and the vacuolar +H-pumping ATPase. The numbers located on the various protein subunits indicate their molecular weight in kilodaltons. The mitochondrial ATP synthase makes ATP from ADP by utilizing the +H-gradient generated by oxidative phosphorylation. The vacuolar ATPase hydrolyzes ATP and pumps +H-ions. F1 and F0 and V1 and V0 complexes are indicated.

The importance of the acidity of the lysosomal lumen is that most of the lysosomal hydrolases are active only at acidic pH. However, certain lysosomal hydrolases such as cathepsin L retain activity even at pH 7.0.1,25 An additional importance for the acidic environment within lysosomes is that many lysosomal enzymes are synthesized as precursors and are properly cleaved to their mature form only at acidic pH.1,7,26 There are many examples of mistargeting of lysosomal or vacuolar proteins when acidification is defective or blocked by specific inhibitors of the V-ATPases such as bafilomycin.27,28 Interpretation of this result is complicated by the fact that many other vesicles and organelles also contain V-ATPases, and their acidity may be required for proper protein targeting. The acidity also undoubtedly helps to denature proteins and other macromolecular substrates to make them easier to digest. The acidification of the yeast vacuole plays additional roles.29,30 The proton gradient and resulting membrane potential drive the transport of amino acids and of ions29 into the vacuole for purposes of nutrient storage and osmotic balance.30 The V-ATPases may be regulated at several different levels. Not all the V-ATPase subunits are assembled into functional pumps in yeast and mammalian cells,31 so assembly factors may regulate pump activity.32 In addition, there is tissue-specific expression of

Introduction

7

isoforms of some subunits.33 The availability of critical sulfhydryl groups is also required for full activity of the proton pump in mammalian cells.34 Finally, the activity of chloride transporter systems in the lysosomal membrane may limit the proton pumping under certain conditions.35

Biogenesis of Mammalian Lysosomes Proteins in the Lysosomal Lumen The different steps in the pathway of synthesis of lysosomes are shown in Figure 1.5. Lysosomal enzymes that are soluble within the lysosomal lumen or matrix are transported to lysosomes due to their specific recognition by receptors for mannose-6-phosphate (M6P).36 Lysosomal enzymes are synthesized like proteins in the secretory pathway (see Chapter 3),7,37,38 but are recognized by a phosphotransferase enzyme shortly after leaving the endoplasmic reticulum (ER). This enzyme transfers N-acetyl-glucosamine-1-phosphate to one or more mannose residues on lysosomal enzymes. A glucosaminidase removes the glucosamine to generate the M6P.7,39 The phosphotransferase recognizes not only the M6P but also polypeptide features within the lysosomal enzymes.7,40 These peptide regions are not linear, but appear to be formed as a result of protein folding.40,41 Nevertheless, progress is being made toward defining the peptide regions that are common among intralysosomal enzymes and are not found in secreted proteins. Lysine residues within a particular distance and in a particular chemical environment may be an essential aspect of the peptide recognition by the phosphotransferase.41,42 The recognition of M6P by the M6P receptor (M6PR) appears to be required for the proper delivery of many lumenal enzymes to lysosomes. Patients with mucolipidosis II (also called I-cell disease) fail to form M6P on lysosomal enzymes due to a mutation in the gene encoding the phosphotransferase which then results in secretion of much of their lysosomal enzymes.1,7,43 There are two types of M6PR in most cells, and both are integral membrane proteins. One is 215 kDa, and the other is 46 kDa.1,7 Figure 1.6 summarizes the structural features of the two M6PRs. The larger receptor binds ligands independently of divalent cations, while the smaller receptor binds proteins that contain M6P more avidly in the presence of divalent cations. These two receptors are often referred to as the cation-independent M6PR (ciM6PR) and the cation-dependent M6PR (cdM6PR), respectively. Both receptors are localized primarily to late endosomes but can also be found in the Golgi and the plasma membrane. It is primarily the ciM6PR that can bind to secreted lysosomal enzymes and mediate their receptor-mediated endocytosis and delivery to lysosomes.44 The late endosomal ciM6PR appears to be responsible for most sorting of newly synthesized lysosomal enzymes to lysosomes, but the cdM6PR also plays a role with some 30% of newly synthesized lysosomal hydrolases.45 The presence of some M6PR on the plasma membrane suggests that missorted lysosomal precursor proteins that are secreted from cells may be reinternalized and delivered to lysosomes.7 This pathway has been confirmed and even used clinically to attempt to replace defective enzyme activities in patients with lysosomal storage disorders.1,46,47 However, in most cells the usual pathway of lysosomal biogenesis appears to occur intracellularly.7,48 M6PR and the recognized lysosomal hydrolases are synthesized on ER-bound polysomes.1,7 They are progressively glycosylated as they travel through the ER and Golgi

8


Fig. 1.5. Pathways for delivering lysosomal matrix proteins to lysosomes. N = nucleus; RER = rough endoplasmic reticulum, G = Golgi; L = lysosome; and PM = plasma membrane. The solid arrows indicate the major route of delivery of lysosomal matrix proteins in most mammalian cells. The open arrows indicate that lysosomal enzymes are also secreted by some cells and then reinternalized by M6PR in the PM.

complex.49 Their mode of movement from the endoplasmic reticulum to the Golgi is within vesicles coated with a protein complex called coatomer that is composed of coat protein (COPII; see Fig. 2.9) molecules and an ADP-ribosylation factor (ARF).50 In the trans-Golgi network (TGN) M6PR and lysosomal proteins are packaged into differently coated vesicles which travel to late endosomes. These vesicles are coated with clathrin heavy and light chain50-55 as are endocytic vesicles responsible for receptor-mediated endocytosis at the plasma membrane (see Chapter 2). The clathrin is recruited to the TGN by adaptor protein-1 (AP-1; see Fig. 2.6) complexes.52 Surprisingly, recruitment of AP-1 to the TGN is due, in part, to the presence of M6PR, a cargo molecule.54 Trafficking of these vesicles also requires an ARF family member.55

Introduction

9

Fig. 1.6. Domain structure of the two M6PR in mammalian cells. The cation independent M6PR (ciM6PR) is a 215 kilodalton integral membrane protein. It consists of a 44-residue aminoterminal ER signal sequence (not shown), a 2269-residue lumenal domain, a single 23-residue transmembrane region, and a 163 amino acid carboxyl terminal cytoplasmic domain. The lumenal domain contains N-linked glycosylation sites (Y) and consists of 15 contiguous repeating segments of approximately 147 amino acids each. The cation dependent M6PR (cdM6PR) is an integral membrane glycoprotein of 46 kilodaltons. Its lumenal domain consists of a single repeat of a sequence similar to the 147 amino acid repeat in the larger receptor. N and C identify amino and carboxyl termini of the proteins.

Interestingly, the ciM6PR at the cell surface also acts as the receptor for insulin-like growth factor II (IGF-II).56 It may be that IGF-II is delivered to lysosomes due to its binding to ciM6PR so that this interaction will keep the circulating half-life of IGF-II short.1,7 Alternatively, binding of IGF-II may alter the normal trafficking of lysosomal hydrolases thereby linking cell growth to altered traffic of lysosomal enzymes.7,57 There are also indications that some lysosomal matrix proteins are targeted to lysosomes by M6P-independent mechanisms. For example, in I-cell disease certain cell types do not

10


accumulate material in lysosomes and appear to contain normal levels of many lysosomal enzymes.1,7,58 Such tissues must have M6P-independent lysosomal targeting mechanisms. Furthermore, no M6PR is evident in yeast,1,7,27,28 and the targeting of lysosomal matrix proteins appears to require a specific peptide signal which, when mutated, results in secretion of the vacuolar protein.59

Proteins in the Lysosomal Membrane Lysosomal membranes contain several abundant glycoproteins (Fig. 1.7) called lysosomal glycoproteins (LGPs) or lysosome-associated membrane proteins (LAMPs) or lysosomal integral membrane proteins (LIMPs).7,60,62 There are several of these proteins. The LAMP-1 and LAMP-2 subtypes have been studied in some detail. They are both highly glycosylated on the lysosomal lumenal side and have a single transmembrane spanning region and a short 11-12 amino acid cytosolic tail.7,60 A conserved tyrosine in the cytosolic tail is important in the correct targeting to the lysosomal membrane, since mutations of this amino acid result in partial mistargeting to the plasma membrane.62 LIMP-I contains the same tyrosine-based lysosomal targeting sequence, while LIMP-II contains a dileucine motif in its cytosolic tail rather than a tyrosine-containing motif.63 Most LAMP-1 and LAMP-2 proteins appear to be targeted to lysosomes by the same route taken for lysosomal matrix proteins except that the M6PR is not involved. The LAMPs are synthesized with an ER signal sequence and appear to travel from the ER to the Golgi and then to late endosomes and finally to lysosomes. A small amount of some forms of LAMP-2 localizes to the plasma membrane and early endosomes.64 Three different splice variants of LAMP-2 exist in avian cells, and these three variants have a common lumenal domain sequence but different transmembrane and cytosolic tail sequences. LAMP2a and LAMP-2b are expressed 17% and 25% at the cell surface, respectively, while LAMP2c is expressed only 6% at the cell surface.65 These studies also implicate a bulky, hydrophobic carboxy-terminal amino acid in addition to the tyrosine for the efficient localization of LAMPs to the lysosomal membrane.65 The functions of LAMP-1 are unknown. Their extensive glycosylation is not required for lysosomal targeting or stability within lysosomes since mutants in terminal glycosylation are not altered in these properties.62 LAMP-2a functions as a receptor for the selective uptake of cytosolic proteins by lysosomes for subsequent degradation (see Chapter 6). LIMP-I and LIMP-II show no homology with the LAMPs. LIMP-I spans the lysosomal membrane four times while LIMP-II spans the lysosomal membrane twice (Fig. 1.7). The functions of LIMP-I and LIMP-II are not known. Lysosomal acid phosphatase (LAP) is synthesized as an integral membrane protein but is cleaved in its final lysosomal residence to become a lysosomal lumenal protein.66 Its synthetic pathway resembles that of the LAMPs and LIMPs. LAP is transported through the endoplasmic reticulum to the trans-Golgi with a T1/2 of 30 min and then appears in lysosomes in its mature form with a T1/2 of 45 hrs. LAP is unrelated to LAMPs or LIMPs, but it also has a short 18 amino acid cytosolic tail, and the tyrosine is an important determinant of proper targeting to lysosomes.67

Biogenesis of the Yeast Vacuole The yeast vacuole is commonly considered to be analogous to lysosomes in mammalian cells. In many respects this view is justified since the yeast vacuole is a hydrolytic organelle,7,68-71 and lysosomes from mammalian cells and the vacuole of yeast can capture intracellular proteins by macroautophagy and microautophagy. However, the yeast vacuole

Introduction

11

Fig. 1.7. Five lysosome-associated membrane proteins (LAMPs). LAMP-1, LAMP-2, LIMP-I, and LIMP-II are integral membrane glycoproteins. Lysosomal acid phosphatase (LAP) is synthesized as an integral membrane protein but is later cleaved so that the mature enzyme resides in the lumen of the lysosome. The protein backbones and glycosylation sites (Y) are indicated. The amino acid sequence of the cytoplasmic tails are indicated in single letter notation. The Y, in the cytosolic tail is required for proper targeting to the lysosomal membrane. Notice that LIMP-II contains a dileucine motif (LI) rather than a Y-based sequence.

serves other important functions that are not shared by mammalian lysosomes. These include storage of inorganic ions, amino acids, and polyphosphates.68 The reader is referred to other recent reviews of these vacuolar functions.72,73 In many respects targeting of vacuolar enzymes to the yeast vacuole resembles targeting of lysosomal enzymes to mammalian lysosomes. For example, most vacuolar proteins are first made as larger precursors with an ER signal sequence.68,71 The precursors are glycosylated in the ER and further glycosylated in the Golgi. Secreted proteins and vacuolar proteins appear to follow the same routes until the late Golgi where vacuolar proteins are then sorted into separate vesicles from those destined for secretion. Vesicles loaded with vacuolar cargo then fuse with the vacuole.68,71 The processing of many vacuolar preproteins requires a functional vacuolar protease A (PrA) gene product, and the normal T1/2 of this processing is 6 min.68,71,74 The vacuolar sorting pathways in yeast do not require M6P or any other glycosylation signal.68,71 A peptide sequence within the vacuolar proteins appears to be necessary and

12


sufficient for vacuolar sorting.68,75 This sequence is commonly near the amino terminus, but little sequence identity is obvious in such vacuolar targeting peptides. There are many variations in this biosynthetic scheme for individual proteins that have been analyzed. For example, carboxypeptidase Y (CPY) is processed in the vacuole in two steps by PrA and then by vacuolar protease B (PrB).68,74 PrB contains both N-linked and O-linked carbohydrates, and a very large propeptide region (281 amino acids) is removed in the ER in addition to further processing by PrA in the vacuole.75 Vacuolar protein sorting (vps) mutants that fail to target CPY to the vacuole fall into 47 different complementation groups.68,71 Class D mutations accumulate many small vesicles in the cytoplasm, and class D mutations are thought to be defective in targeting Golgi-derived vesicles to the vacuole. In fact, many vacuolar proteins can be localized to these small vesicles in the class D mutants. However, another vacuolar membrane protein, alkaline phosphatase (ALP), appears to be targeted to the vacuole through an alternative pathway that is not disrupted by the class D vps mutants (Fig 1.8).76 The entry of ALP into the alternate targeting pathway requires a targeting sequence of 13 amino acids at the cytosolic, carboxyl-terminus of ALP. Finally, another vacuolar membrane protein, Vam3p, is required for this alternative vacuolar targeting pathway to function (Fig. 1.8).71 Two proteins that are soluble within the vacuole appear to be targeted by a different pathway than those described above.68,71,74,78 These proteins, aminopeptidase I and α-mannosidase, are not synthesized with an ER signal sequence and are first localized to the cytosol of yeast. These enzymes are then efficiently delivered to the vacuole by processes that share at least some common elements with pathways of macroautophagy (see Chapter 4).77 The pathway of import of both α−µannosidase and aminopeptidase I is saturable.79,80 A targeting sequence is present in the propeptide region of aminopeptidase I, and this cytosol-to-vacuole targeting pathway has been reproduced using permeablized cells.80 It is not clear why these vacuolar proteins are targeted to vacuoles by a different pathway than most vacuolar proteins. Perhaps these two proteins cannot withstand the conditions within the ER (an oxidizing and high calcium environment). Alternatively, aminopeptidase I and α−µannosidase may require exposure to cytosol prior to delivery to lysosomes for proper posttranslational modification or for multimerization.81

Degradation of Lysosomal Proteins

Resident lysosomal proteins are surprisingly longlived (T1/2 = 5-7 days)82,83 even though they reside in an organelle that has been estimated to contain more than 100 mg/ml of cathepsin D alone.84 Lysosomal enzymes have presumably evolved to have structures that are resistant to the multiple lysosomal proteases and peptidases, but what these structures might be is largely unknown. The glycosylation of lysosomal proteins is often cited as providing a means of protecting the proteins from proteolytic digestion, and glycosylation certainly contributes to the stability of lysosomal proteins.82 Another possible means of protection may be by forming macromolecular complexes. For example, cathepsin A is known to associate with a variety of lysosomal enzymes,85 and in the absence of cathepsin A, these lysosomal proteins become more susceptible to proteolysis.86 A different example comes from studies of the lysosomal molecular chaperone (ly-hsc73). Ly-hsc73 precipitates at pH 5.3 (the protein’s isoelectric point) but remains active within the lysosome. This protein is is also remarkably stable within lysosomes,19 perhaps due to shielding of most of the molecules from proteolytic attack.

Introduction

13

Fig. 1.8. Targeting of vacuolar proteins to the yeast vacuole. At least two different vesicular pathways are able to target proteins from the Golgi to the vacuole. The alternative pathway has been studied both genetically and biochemically, but the characteristics of the vesicles used are largely unknown as indicated by the question mark. Vam3p is another vacuolar membrane protein that travels through the alternative pathway and is also required for the operation of this alternative pathway. Another pathway targets some vacuolar proteins from the cytosol to the vacuole. The proteins that are targeted to the vacuole by this pathway, aminopeptidase I (API) and α-mannosidase, are first concentrated in vesicles that share some properties of macroautophagic vacuoles including the double membrane.

Future Directions of Research There is clearly much left to learn about biogenesis of lysosomes and the yeast vacuole. Continued efforts to identify and characterize the vps mutants in yeast should refine our understanding in this area. In addition, there are important cases of retargeting a lysosomal enzyme for secretion under particular physiological circumstances. For example, many transformed cells secrete cathepsin L while the control nontransformed cells do not.87 A better understanding of such rerouting processes will undoubtedly shed light on new aspects of lysosomal biogenesis. Another area of research that requires more study is the mechanisms by which resident lysosomal enzymes withstand the incredibly harsh conditions within lysosomes. This question may best be addressed using recombinant DNA procedures to create artificial vacuolar targeting sequences on a reporter protein that could then be modified in specific ways. The alterations could be made to change the structure of the reporter protein or to alter its binding to other lysosomal components.

14


Even though regulation of the assembly and activity of the V-ATPase proton pump has been extensively studied, additional work is needed to more completely understand these mechanisms. Acidification is required for the activities of many lysosomal enzymes and is also important for accurate protein trafficking within the cell. Why acidification is important in protein trafficking remains largely unknown. Finally, the roles of LAMP-1, LIMP-I, and LIMP-2 in the lysosomal membrane require further study. These genes could be knocked out in mice and/or their expression could be inhibited by antisense oligonucleotides in cultured cells. Results of such studies must be coupled with regulated expression of the relevant genes since the complete absence of a gene product can cause a variety of indirect compensatory changes. For example, lysosomes may became more fragile in the complete absence of LIMP-I, but expression of a small amount of LIMP-I may correct this fragility. Therefore, while the knock-out mice may exhibit necrotic damage to cells that contain the most lysosomes, graded expression of LIMP-I may show no such effect. References 1. Holtzman E. Lysosomes. New York and London: Plenum Press, 1989:1-439. 2. Pitt D. Lysosomes and Cell Function. London and New York: Longman Group, 1975:1-165. 3. Novikoff AB. Lysosomes in the physiology and pathology of cells: Contributions of staining methods. In: de Reuck AVS and Cameron MP, eds. Lysosomes, Ciba Foundation Symposium. London: Churchill, 1963:36-77. 4. Novikoff AB, Beaufay H, de Duve C. Electron microscopy of lysosome-rich fractions from rat liver. J Biophys Biochem Cytol 1956; 2:179-184. 5. de Duve C, Wattiaux R. Functions of lysosomes. Ann Rev Physiol 1966; 28:435-492. 6. de Duve C. Exploring cells with a centrifuge. Science 1975; 189:186-194. 7. Kornfeld S, Mellman I. Biogenesis of lysosomes. Ann Rev Cell Biol 1989; 5:483-525. 8. Ohkuma S, Poole B. Fluorescence probe measurements of the intralysosomal pH in living cells and the perturbation of pH by various agents. Proc Nat Acad Sci USA 1978; 75:3327-3331. 9. Yamashiro DJ, Maxfield FR. Kinetics of endosome acidification in mutant and wild-type Chinese hamster ovary cells. J Cell Biol 1987; 105:2713-2721. 10. Kim JH, Lingwood CA, Williams DB et al. Dynamic measurement of the pH of the Golgi complex in living cells using retrograde transport of the verotoxin receptor. J Cell Biol 1996; 134:13871399. 11. Diment S, Leech MS, Stahl PD. Cathepsin D is membrane-associated in macrophage endosomes. J Biol Chem 1988; 263:6901-6907. 12. Autheir F, Desbuquois B. Degradation of glucagon in isolated liver endosomes. Biochem J 1991; 280:211-218. 13. Novikoff AB, Novikoff PM. Cytochemical contributions to differentiating GERL from the Golgi apparatus. Histochem J 1977; 9:525-551. 14. Hopkins CR, Gibson A, Shipman M et al. Movement of internalized ligand-receptor complexes along a continuous endosomal reticulum. Nature 1990; 346:335-339. 15. Robinson MS, Watts C, Zerial M. Membrane dynamics in endocytosis. Cell 1996; 84:13-21. 16. Klausner RD. Sorting and traffic in the central vacuolar system. Cell 1989; 57:703-706. 17. Madden EA, Storrie B. Isolation of subcellular organelles. Methods Enzymol 1990; 182:203-225. 18. Mellman I, Fuchs R, Helenius A. Acidification of the endocytic and exocytic pathways. Ann Rev Biochem 1986; 55:663-700. 19. Agarraberes FA, Terlecky SR, Dice JF. An intralysosomal hsp70 is required for a selective pathway of lysosomal protein degradation. J Cell Biol 1997; 137:825-834. 20. Ohkuma S, Chudzik J, Poole B. The effects of basic substances and acidic ionophores on the digestion of exogenous and endogenous proteins in mouse peritoneal macrophages. J Cell Biol 1986; 102:959-966. 21. Forgac M. Structure and properties of the coated vesicle (H+)-ATPase. J Bioenerg Biomembr 1992; 24:341-350.

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22. Pedersen PL, Amzel LM. ATP synthases. Structure, reaction center, mechanism, and regulation of one of nature’s most unique machines. J Biol Chem 1993; 268:9937-9940. 23. Nelson N. Organellar proton-ATPases. Curr Opin Cell Biol 1992; 4:654-660. 24. Forgac M. Regulation of vacuolar acidification. Soc Gen Physiol Ser 1996; 51:121-132. 25. Kirschke H, Langer J, Wiederanders B et al. Cathepsin L. Eur J Biochem 1977; 74:293-301. 26. Hasilik A. The early and late processing of lysosomal enzymes: Proteolysis and compartmentation. Experientia 1992; 48:130-151. 27. Banta LM, Robinson JS, Kionsky DJ et al. Organelle assembly in yeast: characterization of yeast mutants defective in vacuolar biogenesis and protein sorting. J Cell Biol 1988; 107:1369-1383. 28. Klionsky DJ, Emr SD. Membrane protein sorting: Biosynthesis, transport, and processing of yeast vacuolar alkaline phosphatase. EMBO J 1989; 8:2241-2250. 29. Ohsumi Y, Anraku Y. Calcium transport driven by a proton motive force in vacuolar membrane vesicles of Saccharomyces cerevisiae. J Biol Chem 1983; 258:5614-5617. 30. Cramer CL, Davis RH. Polyphosphate-cation interaction in the amino acid-containing vacuole of Neurospora crassa. J Biol Chem 1984; 259:5152-5157. 31. Meyers M, Forgac M. Assembly of the peripheral domain of the bovine vacuolar H(+)-adenosine triphosphatase. J Cell Physiol 1993; 156:35-42. 32. Hill KJ, Stevens TH. Vma22p is a novel endoplasmic reticulum-associated protein required for the asssembly of the yeast vacuolar H+-ATPase complex. J Biol Chem 1995; 270:22329-22336. 33. Forgac M. Structure and function of vacuolar class of ATP-driven proton pumps. Physiol Rev 1989; 69:765-796. 34. Feng Y, Forgac M. A novel mechanism for regulation of vacuolar acidification. J Biol Chem 1992; 267:19769-19772. 35. Mulberg AE, Tulk BM, Forgac M. Modulation of the coated vesicle chloride channel activity and acidification by reversible protein kinase A-dependent phosphorylation. J Biol Chem 1991; 266: 20590-20593. 36. Hoflack B, Kornfeld S. Purification and characterization of a cation-dependent mannose-6-phosphate receptor from murine P388D macrophages and bovine liver. J Biol Chem 1985; 260:1200612014. 37. Ludwig T, Griffiths G, Hoflack B. Distribution of newly synthesized lysosomal enzymes in the endocytic pathway of normal rat kidney cells. J Cell Biol 1991; 115:1561-1572. 38. Rosenfeld MG, Krebich G, Popov D et al. Biosynthesis of lysosomal hydrolases: Their synthesis on bound polysomes and the role of carbohydrate and post-translational processing in determining their subcellular distribution. J Cell Biol 1982; 93:135-143. 39. Kornfeld R, Kornfeld S. Assembly of asparagine-linked oligosaccharides. Ann Rev Biochem 1985; 54:631-664. 40. Reitman ML, Kornfeld S. Lysosomal enzyme, N-acetylglucosaminyl-phosphotransferase, selectively phosphorylates native lysosomal enzymes. J Biol Chem 1981; 256:11977-11980. 41. Lang L, Reitman ML, Tang J et al. Lysosomal enzyme phosphorylation: Recognition of a proteindependent determinant allows specific phosphorylation of oligosaccharides present on lysosomal enzymes. J Biol Chem 1984; 259:14663-14671. 42. Cuozzo JW, Sahagian GG. Lysine is a common determinant for mannose phosphorylation of lysosomal proteins. J Biol Chem 1994; 269:14490-14496. 43. Tager JM. Biosynthesis and deficiency of lysosomal enzymes. Trends Biochem Sci 1985; 10:324-326. 44. Stein M, Zijderhand-Bleekemolen JE, Geuze H et al. Mr 46,000 mannose 6-phosphate-specific receptor: Its role in targeting of lysosomal enzymes. EMBO J 1987; 6:2677-2681. 45. Nolan CM, Creek KE, Grubb J et al. Antibody to the phosphomannosyl receptor inhibits receptor recycling in fibroblasts. J Cell Biochem 1987; 35:137-151. 46. Watts RWE, Gibbs DA. Lysosomal Storage Diseases. London: Taylor and Francis. 1986. 47. Gieselman V. Lysosomal storage diseases. Biochim Biophys Acta 1995; 1270:103-136. 48. Duncan JR, Kornfeld S. Intracellular movement of two mannose 6-phosphate receptors: Return to the Golgi apparatus. J Cell Biol 1988; 106:617-628. 49. Fedde KN, Sly WS. Ricin-binding properties of acid hydrolases from isolated lysosomes implies prior processing by terminal transferases of the trans-Golgi apparatus. Biochem Biophys Res Commun 1985; 133:614-620. 50. Rothman JE. Mechanisms of intracellular protein transport. Nature 1994; 372:55-63. 51. Lemansky P, Hasilik A, von Figura K et al. Lysosomal enzyme precursors in coated vesicles derived from the exocytic and endocytic pathways. J Cell Biol 1987; 104:1743-1748.

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52. Brodsky FM. Living with clathrin: Its role in intracellular membrane traffic. Science 1988; 242:1396-1402. 53. Farquhar MG, Palade GE. The Golgi apparatus: 100 years of progress and controversy. Trends Cell Biol 1998; 8:2-10. 54. Braulke T, Gartung G, Hartung A et al. Is movement of mannose 6-phosphate receptor triggered by binding of lysosomal enzymes? J Cell Biol 1987; 104:1735-1742. 55. Orci L, Palmer DJ, Amherdt M et al. Coated vesicle assembly in the Golgi requires only coatomer and ARF proteins from the cytosol. Nature 1993; 364:732-734. 56. Morgan DO, Edman JC, Standing DN et al. Insulin-like growth factor II receptor as a multifunctional binding protein. Nature 1987; 329:301-307. 57. Kiess W, Thomas CL, Sklar MM et al. Insulin-like growth factor II inhibits both the cellular uptake of β-galactosidase and the binding of β-galactosidase to purified IGF-II/mannose 6-phosphate receptor. J Biol Chem 1989; 264:4710-4714. 58. Neufeld EF, McKusik VA. Disorders of lysosomal enzyme synthesis and localization: I-cell disease and pseudo-Hurler polydystrophy. In: Stanbury JB, Wyngaarden JB, Frederickson DS et al, eds. The Metabolic Basis of Inherited Disease. 5th Edition. New York: McGraw-Hill, 1983:778-787. 59. Valls LA, Hunter CP, Rothman JH et al. Protein sorting in yeast: The localization determinant of yeast vacuolar carboxypeptidase Y resides in the propeptide. Cell 1987; 30:887-897. 60. Howe CL, Granger BL, Hull M et al. Derived protein sequence, oligosaccharides, and membrane insertion of the 120 kDa lysosomal membrane glycoprotein (lgp 120): Identification of a highly conserved family of lysosomal membrane glycoproteins. Proc Nat Acad Sci USA 1988; 85:7577-7581. 61. Fukuda M, Viitala J, Matteson J et al. Cloning of cDNAs encoding human lysosomal membrane glycoproteins, h-lamp-1 and h-lamp-2. Comparison of their deduced amino acid sequences. J Biol Chem 1988; 263:18920-18928. 62. Hunziker W, Geuze H. Intracellular trafficking of lysosomal membrane proteins. BioEssays 1996; 18:379-387. 63. Vega MA, Rodriguez F, Segui B et al. Targeting of lysosomal integral membrane protein LIMP II. J Biol Chem 1991; 266:16269-16272. 64. Uthayakumar S, Granger BL. Cell surface accumulation of overexpressed hamster lysosomal membrane glycoproteins. Cell Mol Biol Res 1996; 41:405-420. 65. Gough NR, Fambrough DM. Different steady state subcellular distribution of the three splice variants of lysosome-associated membrane protein LAMP-2 are determined largely by the COOHterminal amino acid residue. J Cell Biol 1997; 137:1161-1169. 66. Pohlmann R, Crentler C, Schmidt B et al. Human lysosomal acid phosphatase: Cloning, expression, and chromosomal assignment. EMBO J 1988; 7:2343-2350. 67. Prill V, Lehmann L, von Figura K et al. The cytoplasmic tail of lysosomal acid phosphatase contains overlapping but distinct signals for basolateral sorting and rapid internalization in polarized MDCK cells. EMBO J 1993; 12: 2181-2193. 68. Klionsky DJ, Herman PK, Emr SD. The fungal vacuole: Composition, function, and biogenesis. Microbio Rev 1990; 54: 266-292. 69. Klionsky DJ. Protein transport from the cytoplasm to the vacuole. J Membrane Bio 1997; 157:105-115. 70. Klionsky DJ. Nonclassical protein sorting to the yeast vacuole. J Biol Chem 1998; 273:1080710810. 71. Bryant NJ, Stevens TH. Vacuole biogenesis in Saccharomyces cerevisiae: Protein transport pathways to the yeast vacuole. Micro Mol Biol Rev 1998; 62:230-247. 72. Davis RH. Compartmental and regulatory mechanisms in the arginine pathways of Neurospora crassa and Saccharomyces cerevisiae. Microbio Rev 1986; 50:283-313. 73. Anraku Y, Umemoto N, Hirata R et al. Structure and function of the yeast vacuolar membrane proton ATPase. J Bioenerg Biomembr 1989; 21: 589-603. 74. Hemmings BA, Zubenko GS, Hasilik A et al. Mutant defective in processing of an enzyme located in the lysosome-like vacuole of Saccharomyces cerevisiae. Proc Nat Acad Sci USA 1981; 78:435-439. 75. Mechler B, Hirsch H, Muller H et al. Biogenesis of the yeast lysosome (vacuole): Biosynthesis and maturation of yscB. EMBO J 1988; 7:1705-1710. 76. Piper RC, Bryant NJ, Stevens TH. The membrane protein alkaline phosphatase is delivered to the vacuole by a route that is distinct from the VPS-dependent pathway. J Cell Biol 1997; 138:531-545.

Introduction

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77. Harding TM, Hefner-Gravink A, Thumm M et al. Genetic and phenotypic overlap between autophagy and the cytoplasm to vacuole protein targeting pathway. J Biol Chem 1996; 271:17621-17624. 78. Yoshihisa T, Anraku Y. Nucleotide sequence of AMS1, the structural gene of vacuolar α-mannosidase of Saccharomyces cerevisiae. Biochem Biophys Res Commun 1988; 163: 908-915. 79. Klionsky DJ, Cueva-Yaver DS. Aminopeptidase I of Saccharomyces cerevisiae is localized to the vacuole independent of the secretory pathway. J Cell Biol 1992; 119:287-299. 80. Scott SV, Klionsky DJ. In vitro reconstitution of cytoplasm to vacuole protein targeting in yeast. J Cell Biol 1995; 131:1727-1735. 81. Kim J, Scott SV, Oda MN et al. Transport of a large oligomeric protein by the cytoplasm to vacuole protein targeting pathway. J Cell Biol 1997; 137:609-618. 82. Skudlarek MD, Swank RT. Turnover of two lysosomal enzymes in macrophages. J Biol Chem 1981; 256:10137-10144. 83. Hentze M, Hasilik A, von Figure K. Enhanced degradation of cathepsin D synthesized in the presence of the threonine analog β-hydroxynorvaline. Arch Biochem Biophys 1984; 230:372-382. 84. Barrett AJ. Cellular proteolysis. An overview. Ann NY Acad Sci 1992; 674:1-15. 85. Shimmoto M, Fukyhara Y, Ito K et al. Protective protein gene mutations in galactosialidosis. J Clin Invest 1993; 91:2393-2398. 86. Pshezhetsky A, Elsliger M-A, Vinogradova MV et al. Human lysosomal β-galactosidase-cathepsin A complex: Definition of the β-galactosidase-binding interface on cathepsin A. Biochem 1995; 34:2431-2440. 87. Dong J, Sahagian GG. Mechanism for the selective secretion of a lysosomal protease by transformed mouse fibroblasts. J Biol Chem 1989; 264:7377-7383.

CHAPTER 2

Degradation of Endocytosed and Plasma Membrane Proteins Overview

E

ndocytosis refers to a process by which a cell engulfs a particle or a region of the extracellular fluid.1-6 Endocytosis of a particle, such as a bacterium, is called phagocytosis (“cell eating”), while endocytosis of fluid is termed pinocytosis (“cell drinking”). Proteins that enter cells by pinocytosis may do so by fluid-phase, absorptive, and/or receptor-mediated pathways (Fig. 2.1).1-3 Fluid-phase pinocytosis refers to uptake of a compound only by virtue of it being dissolved in the fluid surrounding the cell (Fig. 2.1A).2,4,6 For example, [14C]sucrose is taken up by human fibroblasts in an unsaturable fashion corresponding to the equivalent of 0.31µl/hr/106 cells (Fig. 2.2).4 Uptake of most proteins by most cells is through absorptive pinocytosis which refers to the compound not only being in solution but also interacting weakly with the plasma membrane (Fig. 2.1B). Adsorptive endocytosis is saturable but has a high capacity. The measured uptake of such proteins will exceed that of fluidphase markers by 2- to 100-fold depending on the degree of binding to the plasma membrane (Fig. 2.2). Finally, receptor-mediated endocytosis refers to the uptake of compounds that bind strongly and specifically to receptors on the plasma membrane.3,5 Rates of uptake of such ligands may be 106 to 109 times that of fluid-phase (Figs. 2.1C and 2.2).3,5,6 The mode of endocytosis of an extracellular protein depends on the cell type being studied.6 For example, glucagon enters hepatocytes by receptor-mediated endocytosis since hepatocyes have glucagon receptors in their plasma membrane. However, glucagon enters fibroblasts only by absorptive endocytosis since fibroblasts do not contain glucagon receptors.7 Along with extracellular proteins, certain plasma membrane and vesicular membrane proteins can be delivered to lysosomes through early and late endosomes and subsequently degraded within lysosomes.8,9 Examples of such plasma membrane proteins include the insulin receptor (IR) and epidermal growth factor receptor (EGFR).5,9 Other vesicular proteins delivered to lysosomes for degradation include membrane anchored proteins from the smooth endoplasmic reticulum.5,6 However, the pathway of delivery of such proteins is highly selective, and certain proteins are recycled to the plasma membrane or smooth endoplasmic reticulum (SER) rather than being degraded by lysosomes.9 Additional variations on this theme include the degradation of certain proteins in certain cells within endosomes rather than in lysosomes.10,11 Insulin and glucagon begin


Degradation of Endocytosed and Plasma Membrane Proteins

19

Fig. 2.1. Diagrams showing three different types of endocytosis. A clathrin-coated pit can take up materials in the surrounding fluid by (A) fluid-phase endocytosis, (B) adsorptive endocytosis, and (C) receptor-mediated endocytosis. PM = plasma membrane, ee = early endosome, Y = receptor.

to be degraded within early endosomes of hepatocytes.11 The mechanisms of early endosomal degradation in macrophages involves the production of two forms of cathepsin D. One is targeted to the lysosome lumen, but the other contains a membrane-spanning segment and is targeted to early endosomes.10 Certain proteins avoid digestion altogether and are targeted to another region of the plasma membrane or are secreted from the cell in a process called transcytosis. For example, the polymeric immunoglobulin A receptor is known to be transcytosed from the apical to the basolateral surface of polarized cells.13 Viral proteins and some bacterial toxins can escape from the endosome/lysosome system and enter the cytosol.14,15 One well characterized example is the entry of influenza virus by receptor-mediated endocytosis. When the virus reaches the acidic early endosomes,

20


Fig. 2.2. Biochemical measurements of three different types of endocytosis. Uptake of material by fluidphase is not saturable and corresponds to a certain volume internalized/number of cells/time. Adsorptive endocytosis is weakly saturable, while receptor-mediated endocytosis is readily saturable. In human fibroblasts in culture [14C]sucrose is a fluid-phase marker, while [14C]aldolase is internalized by adsorptive endocytosis, and [14C]insulin-like growth factor I is internalized by receptor-mediated endocytosis. Additional pathways of all forms of endocytosis that do not require clathrin are present in many cells.

a conformational change in the viral coat protein then acts to fuse the lipid bilayer of the virus and the endosome. Certain bacteria alter the early endosome compartment in a mysterious way that prevents its trafficking to late endosomes and lysosomes.16 Many yeast plasma membrane proteins are ubiquitinated as a signal for their delivery to the vacuole for degradation.17-20 Certain receptors in mammalian cells are also ubiquitinated.21-23 There are examples in which the ubiquitination may target the receptor for lysosomal degradation,22 but other examples suggest that some plasma membrane proteins can be ubiquitinated and then degraded by the 26S proteasome rather than by lysosomes.23


21

Methods Used to Study Endocytosis The rate of binding, internalization, and degradation of an extracellular protein can be followed using radioactive forms of the protein.1-3,5,6,24 For example, insulin may be labeled by iodination on tyrosine residues using lactoperoxidase and 125I or 131I. Importantly, this iodination performed under the right conditions leaves the insulin fully active.25 Its binding to cells in culture can be measured at low temperatures where endocytosis is blocked.6,26 All radioactivity associated with the cells after binding at low temperatures can be displaced by a rapid wash at low pH1-3,6,27 or by mild protease treatment6,28 indicating that none of the insulin has been taken into the cell. At 37˚C the radiolabeled insulin binds to the insulin receptors on the plasma membrane, then the radioactivity appears within cells in early and then late endosomes.29 Early and late endosomes can be defined by the presence of particular proteins such as ras-related GTPase 4 (rab4) for early endosomes30 and rab731 or the cation independant mannose-6-phosphate receptor1,32 (ciM6PR) for late endosomes. Early and late endosomes can also be separated by slight density and surface charge differences.33 In most cells the radiolabeled insulin dissociates from the IR in endosomes, but the insulin is degraded to acid-soluble small peptides and amino acids only after the hormone has reached lysosomes.34 Depending upon the cell type, reaching lysosomes may require 20-60 min.2,24,27 However, some cells show degradation of radiolabeled insulin and other polypeptide hormones very rapidly after being endocytosed.11,34 In these cells, including macrophages10 and hepatocytes,11 insulin degradation begins in early endosomes.11,34 Another important technique used in the study of endocytosis has been the subcellular localization of ligands by immunolocalization followed by electron microscopy.3,5,35 ,36 An antibody to the ligand can be localized using a second antibody linked to gold particles of a certain size. Particular subcellular compartments can be identified in the same sections by probing with antibodies for a marker protein of organelles followed by a second antibodygold conjugate. Simultaneous localization of multiple proteins can be achieved if the different second antibodies are linked to different sized gold particles.3,5,35,36 To follow the degradation of plasma membrane receptors, cellular proteins are commonly radiolabeled with an amino acid such as [35S]methionine.6,9,37 The receptor of interest must then be purified from the other radiolabeled cellular proteins, and this is usually accomplished by immunoprecipitation with antibodies to the receptor. Alternatively, the receptor can be purified from detergent-solubilized plasma membrane by affinity chromatography using an immobilized ligand column. For example, the IR can be purified using an insulin-Sepharose column.38 The amount of radioactivity in the purified receptor can then be quantitated at increasing times after cessation of [35S]methionine labeling. An important experimental problem with accurate measurement of protein half-lives is the reutilization of amino acid isotopes after they have been released from proteins during protein degradation. This problem, which leads to artifactually long measured half-lives, can be minimized in cultured cells by adding excess unlabeled amino acids to the culture medium after the labeling.39 The most reliable way to determine whether or not reutilization is occuring is to isolate methiony-ltRNAs from cells at increasing times of chase and to determine whether or not [35S]methionine is present.40,41 In practice, reutilization is quickly eliminated under these conditions. Another way that investigators have attempted to eliminate this reutilization of isotope is to carry out the chase in the presence of inhibitors of protein synthesis.42,43 This approach suffers the drawback that certain pathways of

22


protein degradation require continued protein synthesis to operate.44,45 Therefore, the inhibitor of protein synthesis actually slows the degradation rate of certain proteins. Other ways of studying the endocytosis and degradation of plasma membrane proteins include labeling surface-exposed proteins with 125I catalyzed by a particulate form of lactoperoxidase.37,46,47 The receptor of interest must still be purified from other radioactive proteins, and the structures of some proteins may be damaged by the iodination. However, reutilization of the isotope is not a problem because iodotyrosine cannot be charged on tyrosy-ltRNA.48 Recombinant DNA techniques have been important in many aspects of the study of endocytosis. For example, cDNAs encoding receptors not found in a particular cell type can be transfected into cells to follow its endocytic pathway. These studies can be important because of mechanistic information gained. For example, the transferrin receptor when expressed in different cell types may show different abilities to recycle to the plasma membrane after endocytosis. Detailed biochemical and morphological characterization of these cells can thereby lead to better understanding of receptor recycling. Also, the importance of tyrosine-containing peptides in targeting of proteins to clathrin-coated pits has been extensively analyzed using site-directed mutagenesis to change the tyrosine to other amino acids.49 Another example among many is the use of the yeast two-hybrid system to identify proteins that interact with each other in membrane fusion processes.50 The development of cell-free systems to study various aspects of endocytosis has been pivotal in advancing our understanding in this area. For example, Rothman and his colleagues developed a sensitive way to monitor transport between Golgi stacks (Fig. 2.3).51-53 This assay relies on mutant “donor” cells that have been infected with vaccinia stomatitis virus (VSV) and do not have the Golgi enzyme, N-acetyl-glucosamine transferase-1. The major viral coat glycoprotein is not glycosylated with N-acetyl-glucosamine in these mutant cells. The “acceptor” Golgi are derived from uninfected wild type cells that have an active N-acetyl-glucosamine transferase 1. When the donor and acceptor Golgi fuse, the VSV glycoprotein acquires [3H]N-acetyl-glucosamine from added UDP[3H]N-acetyl-glucosamine. This pathway was shown to involve vesicle budding from the donor Golgi, docking to the acceptor Golgi, and fusion with the acceptor Golgi. Several critical components of this pathway have been identified using this in vitro system (see below). Other cell-free systems have been developed to study vesicular transport from endosomes to lysosomes,54 from the Golgi to the plasma membrane,55 and from the Golgi to late endosomes.56 Cell-free assays for the formation of clathrin-coated pits first used isolated plasma membrane sheets derived from adherent cells that were sonicated to remove most of the other components of the cell.57 Clathrin and adaptor protein-2 (AP-2) complexes could be removed from these plasma membrane sheets by various extractions, and the order and stoichiometry of binding could be established. Current assays can measure coated pit formation and budding of coated vesicles. Such assays demonstrated a role for dynamin and GTP, for example.57,58 A common weakness of these biochemical approaches to the study of endocytosis is that the assays are usually optimized by the researcher, and important intracellular regulators may be missed. Therefore, it is best to also use genetics to analyze endocytosis. In both yeast and mammalian cells selections have been devised that allow for the isolation of mutants defective in endocytosis.59,60 Such endocytosis mutants (end) can be assigned to different complementation groups that affect different stages of endocytosis. For example, certain end mutants fail to form clathrin-coated pits while others fail to form clathrin-


23

Fig. 2.3. Cell-free assay for the fusion of Golgi-derived vesicles. The donor Golgi are derived from cells that are infected with VSV and are making the viral coat protein. The donor cells lack the Golgi enzyme, N-acetylglucosamine transferase-1, so no [3H]N-acetyl-glucosamine is added to the viral coat protein after addition of UDP[3H]Nacetyl-glucosamine. However, when acceptor Golgi are added which contain N-acetyl-glucosamine transferase-1 (GAT), fusion of donor and acceptor elements (arrow) results in radiolabeling of the viral coat protein.

coated vesicles. Others fail to acidify the coated vesicle or are defective in the transport from early endosomes to late endosomes.59,60

Mechanisms of Endocytosis in Mammalian Cells The formation of a membrane invagination followed by a membrane fusion event are common properties of all forms of endocytosis.1-3,5,6,61 The initial steps of phagocytosis and pinocytosis are distinct,1-3,6 but the pathway of delivery of the endocytosed material to lysosomes appears to be very similar.1-3,5,6 The membrane invagination is coated with three clathrin heavy chains and three clathrin light chains arranged into a triskelion and then further into pentagons and hexagons in a complex lattice (Fig. 2.4).58-61 A multi-subunit protein, the adaptor protein-2 complex (AP-2), is responsible for recruiting clathrin to regions of the membrane that are going to form vesicles (Fig. 2.5),58,61,62 while adaptor protein-1 complexes (AP-1) recruit clathrin to the trans-Golgi network (TGN).58 Adaptor protein-3 complexes (AP-3) recruit clathrin and perhaps other coat proteins to endosomal and lysosomal membranes.63,64 The protein composition of these AP complexes is shown in Figure 2.6. These adaptor proteins may be organized by selfassociation within the lipid bilayer or they may bind to other membrane proteins. The assembling clathrin

24


Fig. 2.4. Molecular machinery responsible for receptor-mediated endocytosis. Three clathrin heavy chains and 3 clathrin light chains assemble into triskelions (A), and these further assemble into hexameric and pentameric structures (B) that eventually form a coated pit (C).

structures are believed to provide the energy to form the coated pit since their organization into a lattice is a lower energy state than when the chains are free in solution.58 The adaptor proteins in these various membranes also bind to specific inositol polyphosphates, and such binding inhibits the recruitment of clathrin.65 The AP-2 at the plasma membrane also binds to and concentrates a variety of other membrane proteins into the regions thereby selecting certain membrane proteins to be concentrated within the forming clathrin-coated vesicles. Early results indicated that some plasma membrane proteins were stearically excluded from coated pits,66 but more recent evidence does not support this view. For example, receptors that completely lack a cytosolic tail are not concentrated in coated pits, but neither are they excluded from these structures.58,67


25

Fig. 2.5. Adaptor protein complexes (AP-2) in the plasma membrane (A) bind to clathrin triskelions (B). The further assembly of clathrin structures (C) provides the energy for invagination of the plasma membrane (C).

Certain plasma membrane receptors, including the low density lipoprotein receptor (LDLR) and transferrin receptor (TR), are localized to coated pit regions even in the absence of their ligands while most others, including EGFR, IR, and the two asialoglycoprotein receptors (ARs),5,26 localize in these regions in response to ligand binding. There is growing evidence that the ligand-dependent clustering of the latter type of receptor requires additional cytosolic proteins that may mediate the binding of AP-2 to the receptor.58,68-71 For example, the epidermal growth factor receptor protein substrate of 15 kilodaltons (Eps15) was first described as a substrate for the epidermal growth factor receptor kinase,72 but this protein is localized to all clathrin-coated pits in the plasma membrane by virtue of its binding to AP-2. Eps15 can also stimulate actin polymerization, and may be one link between endocytosis and the cytoskeleton.72 The concentration of membrane receptors into coated pits requires a conserved tyrosine-containing internalization signal in the cytoplasmic domain of the receptors.49 This region of the receptor appears to interact with the AP-2 complex. Examples of such tyrosine-containing internalization signals are: LDLR, FPNPVY; TR, YTRF; AR(major species), YQDL; ciM6PR, YKYSKV; cation-dependent mannose-6-phosphate receptor (cdM6PR), YRGV. These peptide regions can be swapped among these receptors, and they still target the receptors to coated pits.49 Structural analyses indicate that these peptide regions commonly form β-turns.49

26


Fig. 2.6. Comparison of the subunit structures of AP-1, AP-2, and AP-3 complexes. In mammalian cells each AP consists of 4 subunits. The β, µ, and σ family members in AP-1, AP-2, and AP-3 are related by amino acid sequence. The γ, α, and δ chains are unique subunits of AP-1, AP-2, and AP-3, respectively. As mentioned in the text, AP1 and AP-2 recruit clathrin. AP-3 may also recruit clathrin, or it may recruit other vesicular coat proteins. This point is not yet settled.

Another protein called dynamin colocalizes with clathrin and AP-2 in coated pits.58,73,74 Dynamin is a GTPase, and binding of GTP to dynamin causes it to redistribute into a ring at the neck of the coated pit.58,74,75 Upon GTP hydrolysis the dynamin acts to pinch off the pit to form a clathrin-coated vesicle (Fig. 2.7). Interestingly, multiple dynamins exist and may function in somewhat different manners in different tissues.58 The coated vesicle is then rapidly uncoated by the constitutively expressed heat shock cognate protein of 73 kDa (hsc73).58,77-80 The uncoating reaction by hsc73 requires intact clathrin triskelions, clathrin light chains, and hydrolysis of ATP.58,77 Microinjection of anti-hsc73 monoclonal antibodies into living cells inhibits receptor-mediated endocytosis and causes the accumulation of ligands in clathrin-coated structures near the plasma membrane.81 The hsc73 appears to cooperate with the coated vesicle-associated protein, auxilin, which contains a DnaJ motif.58,78 DnaJ is an essential factor for activity of DnaK, an E. coli hsp70 family member. Auxilin binds to hsc73 through the auxilin DnaJ homology domain, and auxilin also contains a clathrin-binding domain.78 When auxilin is present on clathrin-coated vesicles, hsc73 is able to uncoat clathrin cages even without clathrin light chains.58 Whatever the mechanisms of uncoating are in vivo, this uncoated vesicle corresponds to an early endosome.


27

Fig. 2.7. The role of the dynamin GTPase in receptor-mediated endocytosis. Dynamin is concentrated in clathrincoated pits along with clathrin (A). Upon binding GTP, dynamin redistributes to the neck of the clathrin-coated pit (B). Dynamin then pinches off the coated pit coupled to hydrolysis of GTP (C) to form a clathrin-coated vesicle (D).

28


The early endosome compartment consists of mixed vesicular and tubular structures,1,6,82,83 and it is also called the compartment for uncoupling receptor and ligand (CURL).5,6,82 This compartment becomes acidified by the action of a proton-pumping ATPase present in this and other subcellular compartments (see Chapter 1). This acidity appears to dissociate certain ligands from their receptors, and the receptors can then be recycled to the plasma membrane, while the ligand may be delivered to lysosomes for degradation. Other receptors are delivered to lysosomes along with their ligands for degradation.2,3,82 The targeting of certain vesicles back to the plasma membrane for recycling while others are targeted to lysosomes requires specificity in the different vesicles and also in the membrane targets. This specificity is thought to be achieved by proteins called v-SNAREs (soluble N-ethylmaleimide-sensitive protein receptors) in the vesicle and t-SNAREs in the target membranes.51,52,84,85 Several different v-SNAREs and t-SNAREs have been identified.84-87 For example, a v-SNARE, cellubrevin, is involved with vesicular recycling from the CURL back to the plasma membrane.85 Another well-studied model of vesicular traffic involves the regulated exocytosis in nerve terminals.88 In this case the v-SNARE is synaptobrevin and the t-SNARE is a member of the syntaxin protein family.88,89 When the v-SNARE interacts with its target t-SNARE, a membrane-associated rab protein hydrolyzes GTP and docks the vesicle in place.89 The multiple rab proteins on vesicles of different origins provides some of the specificity of vesicular traffic. The complexity of the rab protein family is striking; eleven family members in yeast function at different steps in vesicular movement90 (Table 2.1), and 40 different rabs have been described in mammalian cells.90 Rab5 is important in controlling early endosome fusion events.1,91 While rab4 is required for the recycling of vesicles and cargo from CURL back to the plasma membrane1,92 and rab6 is required for vesicular traffic between the cis and trans-Golgi stacks.1,30 The activities of rab proteins appear to be controlled by other proteins. For example, rab5 GTPase activity is regulated by two other vesicular membrane proteins, rabaptin5 and rabex5.72 After the vesicle has docked at the proper membrane, the vesicle and target membranes must fuse. In order for fusion of lipid bilayers to occur, water must be displaced from the hydrophilic sides of the two membranes.61,93 Membrane fusion is catalyzed by a complex assembly of at least two proteins, NSF (N-ethylmaleimide-sensitive factor) and SNAP (soluble NSF attachment protein) that cycle between membranes and the cytosol. In addition, ATP, GTP, and acyl CoA are required for optimal membrane fusion. The SNAPs bind to both v-SNAREs and t-SNAREs to begin the assembly of the fusion apparatus. The mechanism of fusion is the subject of intense current research effort.1,93-95 A family of other proteins, annexins, play a role in endocytosis and probably other forms of membrane traffic. Annexins are cytosolic proteins that can associate with membranes when their 4 calcium-binding sites are occupied.96 The roles of annexins are unclear, but different family members do associate with different transport vesicles. For example, annexins I and II are preferentially associated with early endosomes while annexins IV and VI are more generally associated with intracellular membranes.96 There is also evidence for nonclathrin-mediated endocytic pathways in cells. For example, a temperature-sensitive mutation in dynamin rapidly stops receptor-mediated endocytosis, but only inhibits fluid-phase pinocytosis by 50%.74 Such nonclathrin-mediated vesicular traffic may be important in controlling the size of the plasma membrane, and it may also be used for the internalization of certain receptors and ligands such as interleukin-2 and its receptor in lymphocytes.85 Clathrin-independent endocytosis has recently been


29

Table 2.1. Rabs in yeast Gene

Transport Step Regulated

YPT1

endoplasmic reticulum to Golgi

SEC4

Golgi to plasma membrane

TPT31 and YPT32

within the Golgi

YPT51, YPT52, and YPT53

plasma membrane to early endosome, and early to late endosome, and Golgi to early endosome

YPT6

late Golgi to endosome and endosome to late Golgi

YPT7

late endosome to vacuole, vacuole to vacuole

YPT 10

?

YPT11

?

For review, see reference 90.

demonstrated in response to cell exposure to sphingomyelinase.97 The extent to which this process occurs in normal cells remains to be established. Calveolae are the plasma membrane structures that are responsible for some nonclathrin-mediated endocytosis. Calveolae are pits in the plasma membrane that are highly enriched for the protein calveolin and for cholesterol, sphingomyelin, lipid-anchored proteins, and protein kinase Cα.98-100 There are multiple calveolin gene products that differ in their properties and in abundance in different cell types. The nonclathrin-mediated uptake of the interleukin-2 receptor mentioned earlier is not by calveolae since these structures do not normally exist in lymphocytes.85 The cytoskeleton is important in many stages of membrane traffic (Fig. 2.8). For example, carrier vesicles that shuttle endocytosed proteins from early to late endosomes move along microtubule tracks.85,101 These vesicles are coated not with clathrin but with the nonclathrin co-atomer protein complex I (COPI)85 originally thought to be associated only with Golgi. The subunit composition of COPI and co-atomer protein complex II (COPII) are compared in Figure 2.9. The transport vesicles between early and late endosomes are linked to microtubules by a protein called CLIP-170 (Fig. 2.8).83,102 This protein can be reversibly phosphorylated, and it binds to microtubules only in its unphosphorylated form.84 More recent results also implicate actin and an unconventional myosin in at least some stages of membrane traffic.85

Degradation of Plasma Membrane Receptors Different receptors in the plasma membrane have different half-lives depending, at least in part, on their rate of delivery to lysosomes for subsequent degradation.46,47,103,104 For example, the LDLR, AR, and TR are efficiently targeted back to the plasma membrane following receptor-mediated endocytosis.5,9,105 For LDLR and AR, this recycling happens after the receptor and ligand have dissociated in the CURL.5 In the case of TR, both the receptor and the ligand recycle after the iron is dissociated from the transferrin in the CURL.5,9 In hepatocytes, less than 1% of the AR internalized is delivered to lysosomes for

30


Fig. 2.8. Roles for the cytoskeleton in endocytosis. Actin filaments are required for all forms of endocytosis in mammalian cells and in yeast. The actin and an unconventional myosin may be required along with clathrin and dynamin for early endosome formation. Vesicular traffic between early endosomes (ee) and late endosomes (le) and between le and lysosomes is directed along microtubule tracks in the direction indicated by the solid arrows. PM = plasma membrane; L = lysosome; G = Golgi.

degradation. 37,103 In other words, the AR can interact with more than 100 asialoglycoprotein molecules before the AR is itself degraded. The half-lives of such receptors is between 20 and 60 hrs (see Table 2.2).103 Two examples of plasma membrane receptors that are not efficiently recycled, at least in certain cell types, are the IR and the EGFR. Both of these receptors have half-lives on the order of 8-10 hrs.106-110 Whether or not the degradation rate of the receptor is increased in response to ligand binding depends critically on the cell type studied. For example, IR is degraded more rapidly in the presence of insulin in mouse 3T3 fibroblasts104 and in human lymphocytes106 but not in chick hepatocytes.109 The EGFR half-life varies in different cell types from 10-24 hrs in the absence of ligand to 1-12 hrs in the presence of ligand.37,107,108,110 This increased degradation of EGFR in the presence of


31

Fig. 2.9. Subunit structure of COPI and COPII vesicular coats. (A) COPI subunit sizes in kilodaltons (kDa) are from mammals, but functional homologs also exist in yeast. The interactions between the different subunits are largely unknown. (B) COPII subunit structure was first elucidated in yeast, but homologs also exist in mammalian cells. Mutants in any subunit of COPII are defective in protein secretion (sec). Mutations in sar1, a GTPase, are pleiotropic but are also defective in protein secretion. The molecular weights listed in kDa are from yeast. A cytosolic complex of sec23p-sec24p as well as cytoplasmic sar1p bind to the integral endoplasmic reticulum (ER) membrane protein, sec12p. A cytoplasmic complex of sec13p-sec31p is recruited to the ER membrane due to interactions with the ER membrane protein, sec16p. All of the subunits then interact to form COPII.

epidermal growth factor may account, in part, for the decreased sensitivity of cells to the ligand in response to continuous exposure.107,108,110 Half-lives of other plasma membrane proteins are heterogeneous in hepatocytes.9,37,47,104 Some proteins have half-lives >100 hrs (see Table 2.2), and such long half-


32

Table 2.2. Half-lives of plasma membrane proteins in hepatocytes Protein

Half-Life (hr)

Reference

gap junction protein

4-5

119

insulin receptor

10

106

Na /K -ATPase

18

120

asialoglycoprotein receptor

18

9

nucleotide pyrophosphatase

24

121

ectoATPase

72

122

clathrin heavy chain

>100

47

actin

>100

47

120

122

+

+

dipeptidyl peptidase IV

lives may reflect exclusion from the endocytic machinery and/or efficient recycling to the cell surface.37,47 Proteins with half-lives shorter than 100 hrs are probably included in the endocytic pathway and/or are less efficiently recycled.37,47 This heterogeneity in plasma membrane protein half-lives is not unique to hepatocytes since similar heterogeneity is evident for plasma membrane proteins from fibroblasts (Table 2.3). Another possible fate of plasma membrane proteins is to be ubiquitinated and then degraded by the 26S proteasome. This pathway has been shown for two plasma membrane proteins, the gap junction protein, connexin 43,23 and the heregulin receptor, ErbB-4.111 In the latter example, the ErbB-4 is first cleaved by a metalloprotease active at the cell surface. Only in this cleaved form is the ErbB-4 a substrate for the ubiquitin/26S proteasome pathway of proteolysis.111 Degradation of other integral membrane proteins in the endoplasmic reticulum (ER)112 and even for lumenal proteins in the ER113,114 has also been shown to be by the ubiquitin/proteasome machinery. Since the 26S proteasome is able to digest integral membrane proteins completely, the proteasome must be able to pull the substrate proteins out of the membrane bilayer.113,114 Ubiquitination of some plasma membrane receptors in mammalian cells can target their degradation by lysosomes rather than by the 26S proteasome.22,115 Still other receptors are ubiquitinated, but this modification does not affect the receptor’s degradation rate.21

Receptor-Mediated Endocytosis in Yeast Conjugation of ubiquitin to certain yeast plasma membrane proteins targets the protein for vacuolar degradation.17-20 In the case of the α mating factor receptor, phosphorylation of the protein precedes its ubiquitination.116 Another yeast plasma membrane protein, uracil permease, is tagged with ubiquitins that are linked through lysine 63 rather than the more usual lysine 48.19 Finally, a broad specificity amino acid transporter in the yeast plasma membrane is ubiquitinated and degraded within the vacuole.20 In yeast as in mammals deletion of clathrin heavy chain genes reduces but does not completely block endocytosis.58 However, other aspects of receptor-mediated endocytosis in yeast appear to differ from the process in mammalian cells. There is no evidence for recycling of receptors to the plasma membrane in yeast.58,72 In yeast there is no role for


33

Table 2.3. Half-lives of plasma membrane proteins in fibroblasts Protein

Half-Life (hr)

Reference

Platelet-derived growth factor receptor

1

123

ErbB-4 receptor (80 kDa)

3

111

epidermal growth factor receptor

10

110

glucose transporter

15

124

P-glycoprotein

15

125

c-ras

20

126

low density lipoprotein receptor

25

26

mannose-6-phosphate receptor (cation independent)

26

127

glucocerebrosidase

39

128

dynamin-related proteins in the endocytosis of at least one plasma membrane protein, the α mating factor receptor.58,72 Two dynamin-related proteins, based on their sequence similarities, exist in yeast, Vps1p and Dnm1p, but have distinct roles in targeting of proteins to the vacuole and in endosomal movement, respectively.117,118 A role for Vps1 p in the formation of clathrin-coated vesicles derived from the Golgi is possible.

Future Directions of Research Further definition of the proteins, lipids, and other molecules that are required for endocytosis are necessary for a complete understanding of this process. The challenge in these experiments is to obtain both biochemical and genetic evidence for conclusions that are drawn. The biochemical approach alone may reveal a required factor only under the artificial conditions employed. On the other hand, genetic evidence alone will not distinguish between primary and secondary effects of the mutation. Figure 2.10 indicates the probable complexity of endocytosis when these mechanistic studies have been completed. The molecules in Figure 2.10 that have not been described in this chapter (stabilin, disruptin, amphiphysin, heterobrevin, and homobrevin) are fictitious, but serve to emphasize that we have not yet discovered all the mechanistic features of endocytosis. The roles of lipids in membranes are just beginning to be appreciated and will require much further study. For example, plasma membranes and endocytic membranes contain more phosphatidylserine, sphingomyelin, glycolipids, and sterols than other intracellular membranes.72 Phosphatidylserine on the cytoplasmic face of the plasma membrane may activate specific proteins. There is also evidence that different membrane proteins and lipids may assemble into separate “rafts” within the plasma membrane.72 The pathway of degradation of additional receptors and other plasma membrane proteins need to be documented since clear examples of degradation by lysosomes and by

34


Fig. 2.10. Identified and hypothetical requirements for receptor-mediated endocytosis in mammalian cells. In addition to AP-2, clathrin, receptors, and dynamin, there is also evidence implicating other proteins in receptormediated endocytosis in certain cell types. These molecules include annexins, synaptojanin, Eps15, Eps15 kinase and Eps15 phosphatase (see the text for description). Additional proteins that may prove to play a role in receptormediated endocytosis are fictional at this time. Such proteins include stabilin, disruptin, amphiphysin, homobrevin, and heterobrevin, and they are included in this Figure to emphasize that not all components of the molecular machinery responsible for receptor-mediated endocytosis have been identified.

the ubiquitin/26S proteasome already exist. Ideas about the importance of being degraded by either of the two pathways may be addressed once we know the molecular signals that target receptors to lysosomes and to the 26S proteasome. For example, if EGFR could be slightly modified so that it could be degraded by the 26S proteasome rather than by lysosomes, experiments may show that EGFR can no longer be down regulated in response to prolonged exposure to EGF. References 1. Alberts B, Bray D, Lewis J et al. Molecular Biology of the Cell. 3rd ed. New York and London: Garland Publishing, 1994:551-651. 2. Steinman RM, Mellman IS, Muller WA et al. Endocytosis and recycling of plasma membrane. J Cell Biol 1983; 96:1-27. 3. Stahl P, Schwartz AL. Receptor-mediated endocytosis. J Clin Invest 1986; 77:657-662. 4. Gurley R, Dice JF. Degradation of endocytosed proteins is unaltered in senescent fibroblasts. Cell Biol Internat Rep 1988; 12:885-894. 5. Bu G, Schwartz AL. Receptor-mediated endocytosis. In: Arias I, ed. The Liver: Biology and Pathobiology. 3rd ed. New York: Raven Press, 1994:259-274. 6. Besterman JM, Low RB. Endocytosis: A review of mechanisms and plasma membrane dynamics. Biochem J 1983; 210:1-13. 7. Porterfield SP. Endocrine Physiology. St. Louis: Mosby 1996:105-126. 8. Baldwin DB, Prince M, Tsai P et al. Insulin binding, internalization, and receptor regulation in cultured human fibroblasts. Am J Physiol 1981; 241:E251-E260. 9. Warren R, Doyle D. Turnover of the surface proteins and the receptor for serum asialoglycoproteins in primary cultures of rat hepatocytes. J Biol Chem 1981; 256:1346-1355. 10. Diment S, Leech MS, Stahl PD. Cathepsin D is membrane-associated in macrophage endosomes. J Biol Chem 1988; 263:6901-6907. 11. Autheir F, Desbuquois B. Degradation of glucagon in isolated liver endosomes. Biochem J 1991; 280:211-218.


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42. Fulks RM, Li JB, Goldberg AL. Effects of insulin, glucose, and amino acids on protein turnover in rat diaphragm. J Biol Chem 1975; 250:290-298. 43. Russell DH, Snyder SH. Amine synthesis in regenerating rat liver: Extremely rapid turnover of ornithine decarboxylase. Mol Pharm 1969; 5:253-262. 44. Chiang H-L, Schekman R. Regulated import and degradation of a cytosolic protein in the yeast vacuole. Nature 1991; 350:313-318. 45. Amenta JS, Brocher SC. Mechanisms of protein turnover in cultured cells. Life Sci 1981; 28: 1195-1208. 46. Tweto J, Doyle D. Turnover of the plasma membrane proteins of hepatoma tissue culture cells. J Biol Chem 1976; 251:872-882. 47. Chu FF, Doyle D. Turnover of plasma membrane proteins in rat hepatoma cells and primary cultures of rat hepatocytes. J Biol Chem 1985; 260:3097-3107. 48. Neff NT, Bourret E, Miao P et al. Degradation of proteins microinjected into IMR-90 human diploid fibroblasts. J Cell Biol 1981; 91:184-194. 49. Trowbridge I, Collawn JF, Hopkins CR. Signal-dependent membrane protein trafficking in the endocytic pathway. Annu Rev Cell Biol 1993; 9:129-161. 50. Fields S, Sternglanz R. The two-hybrid system: An assay for protein-protein interactions. Trends Gen 1994; 10:286-292. 51. Rothman JE. The reconstitution of intracellular protein transport in cell-free systems. Harvey Lect 1992; 86:65-85. 52. Beckers JM, Rothman JE. Transport between Golgi cisterna. Methods Enzymol 1992; 219:5-12. 53. Balch WE, Dunphy WG, Braell WA et al. Reconstitution of the transport of proteins between successive compartments of the Golgi measured by the coupled incorporation of N-acetylglucosamine. Cell 1984; 39:405-416. 54. Mullock BM, Bright NA, Fearon CW et al. Fusion of lysosomes with late endosomes produces a hybrid organelle of intermediate density and is NSF-dependent. J. Cell Biol 1998; 140:591-601. 55. Pryer NK, Westhube LJ, Schekman R. Vesicle-mediated protein sorting. Annu Rev Biochem 1992; 61:471-516. 56. Gruenberg J, Clagur MJ. Regulation of intracellular membrane transport. Curr Opin Cell Biol 1992; 5:636-640. 57. Schmid SL. Biochemical requirements for the formation of clathrin- and COP-coated transport vesicles. Curr Opin Cell Biol 1993; 5:621-627. 58. Schmid SL. Clathrin-coated vesicle formation and protein sorting: An integrated process. Annu Rev Biochem 1997; 66:511-548. 59. Raths S, Rohrer J, Crausaz F et al. end3 and end4: Two mutants defective in receptor-mediated endocytosis and fluid-phase endocytosis in Saccharomyces cerevisiae. J Cell Biol 1993; 120:55-65. 60. Robbins AR, Oliver C, Bateman JL et al. A single mutation in Chinese hamster ovary cells impairs both Golgi and endosomal function. J Cell Biol 1984; 99:1296-1308. 61. White JM. Membrane fusion. Science 1992; 258:917-923. 62. Janin J, Chothia C. The structure of protein-protein recognition sites. J Biol Chem 1990; 265:16027-16030. 63. Stepp JD, Huang L, Lemmon SK. The yeast adaptor protein complex, AP-3, is essential for the efficient delivery of alkaline phosphatase to the vacuole. J Cell Biol 1997; 139:1761-1774. 64. Odorizzi G, Cowles CR, Emr SD. The AP-3 complex: A coat of many colours. Trends Cell Biol 1998; 8:282-288. 65. de Camilli P, Emr SD, McPherson PS et al. Phosphoinositides as regulators of membrane traffic. Science 1996; 271:1533-1539. 66. Bretscher MS, Thomson JN, Pearse BMF. Coated pits act as molecular filters. Proc Nat Acad Sci USA 1980; 77:4156-4159. 67. Vigers GPA, Crowther RA, Pearse BMF. Location of the 100kd-50kd accessory proteins in clathrin coats. EMBO J 1986; 5:2079-2085. 68. Pelchen-Matthews A, Boulet I, Littman DR et al. The protein kinase p56lck inhibits CD4 endocytosis by preventing entry of CD4 into coated pits. J Cell Biol 1992; 117:279-290. 69. West MA, Bretscher MS, Watts C. Distinct endocytic pathways in epidermal growth factor-stimulated human carcinoma A431 cells. J Cell Biol 1989; 110:2731-2769. 70. Carpentier JL, McClain D. Insulin receptor kinase activation releases a constraint maintaining the receptor on microvilli. J Biol Chem 1995; 270:5001-5006.


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71. Collawn JF, Stangel M, Kuhn LA et al. Transferrin receptor internalization sequence YXRF implicates a tight turn as the structural recognition motif for endocytosis. Cell 1990; 63:1061-1072. 72. Riezman H, Woodman PG, van Meer G et al. Molecular mechanisms of endocytosis. Cell 1997; 91:731-738. 73. McClure SJ, Robinson PJ. Dynamin, endocytosis, and intracellular signaling. Mol Membr Biol 1996; 13:189-215. 74. Warnock DE, Schmid SL. Dynamin GTPase, a force-generating molecular switch. BioEssays 1996; 18:885-893. 75. Sweitzer SM, Hinshaw JE. Dynamin undergoes a GTP-dependent conformational change causing vesiculation. Cell 1998 93:1021-1029. 76. Jones SM, Howell KE, Henley JR et al. Role of dynamin in the formation of transport vesicles from the trans Golgi network. Science 1998; 279:573-577. 77. Schmid SL, Rothman JE. Enzymatic dissociation of clathrin cages in a two-stage process. J Biol Chem 1985; 260:10050-10056. 78. Holstein SE, Ungewickell H, Ungewickell E. Mechanism of clathrin basket dissociation: Separate functions of the DnaJ homologue, auxilin. J Cell Biol 1996; 135: 925-937. 79. Gao B, Emoto Y, Greene L et al. Nucleotide binding properties of bovine brain uncoating ATP-ase. J Biol Chem 1993; 268:8507-8514. 80. Chappell TG, Welch WJ, Schlossman DM et al. Uncoating ATPase is a member of the 70 kilodalton family of stress proteins. Cell 1986; 45:3-11. 81. Honing S, Kreimer G, Robenek H et al. Receptor-mediated endocytosis is sensitive to antibodies against the uncoating ATPase (hsc70). J Cell Sci 1994; 107:1185-1196. 82. Helenius A, Mellman I, Wall D et al. Endosomes. Trends Biochem Sci 1983; 8:245-250. 83. Parton RG, Schrotz P, Bucci C et al. Plasticity of early endosomes. J Cell Sci 1992; 103: 335-348. 84. Waters MG, Griff LC, Rothman JE. Proteins involved in vesicular transport and membrane fusion. Curr Opin Cell Biol 1991; 3:615-620. 85. Robinson MS, Watts C, Zerial M. Membrane dynamics in endocytosis. Cell 1996; 84: 13-21. 86. Advani RJ, Bae H-R, Bock JB et al. Seven novel mammalian SNARE proteins localize to distinct membrane compartments. J Biol Chem 1998; 273:10317-10324. 87. Rowe T, Dascher C, Bannykh S et al. Role of vesicle-associated syntaxin 5 in the assembly of preGolgi intermediates. Science 1998; 279:696-700. 88. de Camilli P, Takei K. Molecular mechanisms in synaptic vesicle endocytosis and recycling. Neuron 1996; 16: 481-486. 89. Sollner T, Whiteheart SW, Brunner M et al. SNAP receptors implicated in vesicle targeting and fusion. Nature 1993; 362:318-324. 90. Lazar T, Götte M, Gallwitz D. Vesicular transport: How many Ypt/Rab GTPases make a eukaryotic cell? Trends Biochem Sci 1997; 264:468-472. 91. Zerial M, Stenmark H. Rab GTPases in vesicular transport. Curr Opin Cell Biol 1993; 5:613-620. 92. Marsh M, Cutler D. Membrane traffic: Taking the Rabs off endocytosis. Curr Biol 1993; 3:30-33. 93. Zimmerberg J, Vogel SS, Chernomordik IV. Mechanisms of membrane fusion. Annu Rev Biophys Biomol Struct 1993; 22: 433-466. 94. Calakos N, Bennet M, Petersen K et al. Protein-protein interactions contributing to the specificity of intracellular vesicular trafficking. Science 1994; 263:1146-1149. 95. Weber T, Zemelman BV, McNew JA et al. SNARE pins: Minimal machinery for membrane fusion. Cell 1998; 92:759-772. 96. Gruenberg J, Emans N. Annexins in membrane traffic. Trends Cell Biol 1993; 3:224-227. 97. Zha X, Pierini LM, Leopold PL et al. Sphingomyelinase treatment induces ATP-independent endocytosis. J Cell Biol 1998; 140:39-47. 98. Andersen RGW. Potocytosis of small molecules and ions by caveolae. Trends Cell Biol 1993; 3:69-72. 99. Parton RG. Caveolae and caveolins. Curr Opin Cell Biol 1996; 8:542-548. 100. Mineo C, Ying YS, Chapline C et al. Targeting of protein kinase Cα to caveolae. J Cell Biol 1998; 141:601-610. 101. Bloom GS, Goldstein LSB. Cruising along microtubule highways: How membranes move through the secretory pathway. J Cell Biol 1998; 140:1277-1280. 102. Pierre P, Scheel J, Rickard et al. CLIP-170 links endocytic vesicles to microtubules. Cell 1992; 70:887-900.

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103. Baumann H, Hou E, Doyle D. Insertion of biologically active membrane proteins from the rat liver into the plasma membrane of mouse fibroblasts. J Biol Chem 1980; 255:10001-10017. 104. Hare JF. Mechanisms of membrane protein turnover. Biochim Biophys Acta 1990; 1031:71-90. 105. Omary MB, Trowbridge IS. Biosynthesis of the human transferrin receptor in cultured cells. Insulin-induced receptor loss in cultured human lymphocytes is due to accelerated receptor degradation. J Biol Chem 1981; 256:12888-12892. 106. Kasuga M, Kahn CR, Hedo JA et al. Biosynthesis of the human transferrin receptor in cultured cells. Insulin-induced receptor loss in cultured human lymphocytes is due to accelerated receptor degradation. J Biol Chem 1981; 256:6917-6921. 107. Decker SJ. Effects of epidermal growth factor and 12-o-tetradecoylphorbol-13-acetate on metabolism of the epidermal growth factor receptor in normal human fibroblasts. Mol Cell Biol 1984; 4:1718-1724. 108. Krupp MN, Connolly DT, Lane MD. Synthesis, turnover, and down regulation of epidermal growth factor receptors in human A431 epidermoid carcinoma cells and skin fibroblasts. J Biol Chem 1982; 257:11489-11496. 109. Krupp MN, Lane MD. Evidence for different pathways for the degradation of insulin and insulin receptor in the chick liver cell. J Biol Chem 1982; 257: 1372-1377. 110. Stoscheck CM, Carpenter G. Characterization of the metabolic turnover of epidermal growth factor receptor protein in A-431 cells. J Cell Physiol 1984; 120:296-302. 111. Vecchi M, Carpenter G. Constitutive proteolysis of the ErbB-4 receptor tyrosine kinase by a unique, sequential mechanism. J Cell Biol 1997; 139:995-1003. 112. Ward CL, Omura S, Kopito RR. Degradation of CFTR by the ubiquitin-proteasome pathway. Cell 1995; 83:121-127. 113. Hiller MM, Finger A, Schweiger M et al. ER degradation of a misfolded lumenal protein by the cytosolic ubiquitin-proteasome pathway. Science 1996; 273:1725-1728. 114. Plemper RK, Böhmler S, Bordallo J et al. Mutant analysis links the translocon and BIP to retrograde protein transport for ER degradation. Nature 1997; 388:891-895. 115. Strous GJ, van Kerkhof P, Govers R et al. The ubiquitin conjugation system is required for ligandinduced endocytosis and degradation of the growth hormone receptor. EMBO J 1996; 14:3806-3812. 116. Hicke L, Zanolari B, Riezman H. Cytoplasmic tail phosphorylation of the α factor receptor is required for its ubiquitination and internalization. J Cell Biol 1998; 141:349-358. 117. Vater CA, Raymond CK, Ekena K et al. The VPS1 protein, a homolog of dynamin required for vacuolar protein sorting in Saccharomyces cerevisiae, is a GTPase with two functionally separable domains. J Cell Biol 1992; 119:773-786. 118. Gammie AE, Kurihara KL, Vallee RB et al. DNM1, a dynamin-related gene, participates in endosomal trafficking in yeast. J Cell Biol 1995; 130:553-566. 119. Fallon RF, Goodenough DA. Five-hour half-life of mouse liver gap junction protein. J Cell Biol 1981; 90:521-526. 120. Karin NJ, Cook JS. Turnover of the catalytic subunit of the Na,K-ATPase in HTC cells. J Biol Chem 1986; 261:10422-10428. 121. Elovson J. Biogenesis of plasma membrane glycoproteins: Tracer kinetic study of two rat liver plasma membrane glycoproteins in vivo. J Biol Chem 1980; 255:5816-5825. 122. Scott LJ, Hubbard AL. Dynamics of four rat liver plasma membrane proteins and polymeric IgA receptor: Rates of synthesis and selective loss into the bile. J Biol Chem 1992; 267:6099-6106. 123. Claesson-Welsh L, Ronnstrandt L, Heldin C. Biosynthesis and intracellular transport of the receptor for platelet-derived growth factor. Proc Nat Acad Sci USA 1987; 84:8796-8800. 124. White MK, Weber MJ. Transformation by the src oncogene alters glucose transport into rat and chicken cells by different mechanisms. Mol Cell Biol 1988; 8:138-144. 125. Muller C, Laurent G, Ling V. P-glycoprotein stability is affected by serum deprivation and high cell density in multidrug-resistant cells. J Cell Physiol 1995; 163:538-544. 126. Ulsh LS, Shih TY. Metabolic turnover of human c-rasH p21 protein of EJ bladder carcinoma and its normal cellular and viral homologs. Mol Cell Biol 1984; 4:1647-1652. 127. Creek KE, Sly WS. Biosynthesis and turnover of the phosphomannosyl receptor in human fibroblasts. Biochem J 1983; 214:353-360. 128. Das P, Murray GJ, Barranger JA. Studies on the turnover of glucocerebrosidase in cultured rat peritoneal macrophages and normal human fibroblasts. Eur J Biochem 1986; 154:445-450.

CHAPTER 3

Lysosomal Degradation of Proteins in the Secretory Pathway: Crinophagy Overview

M

any cells secrete proteins constitutively. That is, there is a relatively constant flow of proteins through the endoplasmic reticulum (ER) and Golgi complex and then to the plasma membrane where carrier vesicles fuse with the plasma membrane and release their contents outside the cell (Fig. 3.1).1,2 Such cells include fibroblasts, where the secreted proteins are largely components of the extracellular matrix, and hepatocytes, where the secreted proteins circulate in the serum. Other cells exhibit regulated secretion in which the secretory proteins are packaged after reaching the trans-Golgi network (TGN) into secretory vesicles that remain in the cytoplasm until a secretory signal is received (Fig. 3.2).1-3 These cells include a wide variety of endocrine and exocrine cells as well as neurons. Many secreted proteins are made as larger precursor molecules. They are cleaved to the mature protein within the secretory vesicle. Proteins traversing both the constitutive and regulated secretory pathways may be diverted to lysosomes for degradation (Fig. 3.3).4,5 This process, called crinophagy,4 requires vesicular fusion with lysosomes rather than with the plasma membrane. The percentage of secretory proteins delivered to lysosomes for degradation often increases as demand for the secreted product diminishes.4-6

Methods Used to Study Protein Secretion and Crinophagy The major intracellular pathway for regulated protein secretion was elucidated by Palade and coworkers.7 They studied cells of the exocrine pancreas which produce a variety of hydrolases such as trypsin, chymotrypsin, deoxyribonuclease, and ribonuclease. These proteins represent most (>80%) of newly synthesized proteins in these cells, so their intracellular localization could be followed simply by tracking the newly synthesized proteins after short exposure to radioactive amino acids and increasing periods of chase.7 The location of the newly synthesized proteins was followed both by cell fractionation and by microscopy and autoradiography. Both experimental approaches showed that the secreted hydrolases were first synthesized on ribosomes bound to the ER. They were then glycosylated within the ER and further glycosylated within the Golgi. From the Golgi, the hydrolases were packaged into secretory granules called zymogen granules in the exocrine pancreas. The zymogen granules were localized close to the plasma membrane and could be stored in the cell for long periods. These zymogen granules contained all of the hydrolases concentrated more than 100-fold above concentrations in the ER lumen. This concentration Lysosomal Pathways of Protein Degradation, by J. Fred Dice, Ph.D. ©2000 EUREKAH.COM

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Fig. 3.1. Diagram of a cell showing the constitutive secretion pathway. N = nucleus; RER = rough endoplasmic reticulum; G = Golgi; PM = plasma membrane. The constitutive secretory vesicles are the small vesicles shown between the Golgi and the plasma membrane. Arrows indicate the progressive direction of movement of secreted proteins.

began in the Golgi and continued as the zymogen granules matured. After appropriate stimuli associated with food entering the intestine, the secretory granules fused with the plasma membrane of the exocrine cell to release the hydrolases into the intestine.7 More recent studies of protein secretion commonly use antibodies to the protein linked to electron-dense gold particles and follow the intracellular location of the secreted protein by electron microscopy.8,9 In such studies the synthesis of the protein of interest should be capable of being regulated so that a time course of intracellular localization can be followed after turning on synthesis. Alternatively, biochemical analysis may employ

Lysosomal Degradation of Proteins in the Secretory Pathway: Crinophagy

41

Fig. 3.2. Diagram of a cell showing both constitutive and regulated secretory pathways. N = nucleus; RER = rough endoplasmic reticulum; G = Golgi; PM = plasma membrane; SV = secretory vesicles in the regulated secretory pathway. The constitutive secretory vesicles are the small vesicles shown between the Golgi and the plasma membrane. Arrows indicate the direction of movement of secreted proteins.

antibodies to a particular secreted protein along with pulse-chase and subcellular fractionation to follow its subcellular localization at different times after synthesis.10 Such studies have shown that particular proteins may be packaged into secretory granules but also may be secreted by the constitutive pathway in the same cells.11 As in all areas of cell biology, recombinant DNA approaches have been important in our understanding of protein secretion pathways. For example, molecular biology has been essential for defining the peptide signals within proteins that target them for packaging

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Fig. 3.3. Diagrammatic representation of crinophagy. N = nucleus; RER = rough endoplasmic reticulum; G = Golgi; PM = plasma membrane; SV = secretory vesicles in the regulated secretory pathway; L = lysosome. The constitutive secretory vesicles are the small vesicles shown between the Golgi and the plasma membrane. Arrows indicate the direction of movement of secreted proteins and the fact that both constitutive and regulated secretory vesicles may fuse with lysosomes rather than with the plasma membrane.

into secretory granules12 and in establishing the constitutive secretory pathway as the “default” pathway traveled by proteins that enter the ER but have no additional targeting signals.13 In addition, different cell types can be transfected with cDNAs encoding a particular regulated secretory protein. Such studies have shown that most, but not all, cells contain the machinery for packaging of such proteins into secretory granules and for proper proteolytic maturation of the secretory protein.14,15


43

The development of cell-free systems for the study of protein secretion has greatly advanced our understanding of the biochemical requirements for both regulated and constitutive secretory pathways.16-18 Requirements for cytosol and hydrolyzable ATP are common in a variety of these cell-free systems. Factors required for fusion of secretory granules with the plasma membrane include the N-ethylmaleimide-sensitive factor (NSF), soluble NSF attachment proteins (SNAPs), vesicle NSF attachment receptors (v-SNAREs), target membrane NSF attachment receptors (t-SNAREs), and ADP-dependent ribosylation factor (ARF).16,18 Annexin family members are also required and may, in part, account for the calcium-sensitivity of regulated exocytosis.19 Yeast have been especially important in our understanding of mechanisms of constitutive protein secretion. There are now more than 40 different complementation groups of secretion mutants (sec) in S. cerevisiae, and many of the gene products have been identified and studied biochemically.20,21 These genetic studies have emphasized the similarities in mechanisms of protein secretion in yeast and animal cells; in many cases a yeast mutant can be rescued by a mammalian homolog of the mutated gene.22 The study of crinophagy has relied on a combination of morphological and biochemical approaches. The fusion of secretory vesicles with lysosomes has been observed in many tissues that exhibit regulated secretion (Fig. 3.3).4,23,24 However, a portion of proteins undergoing constitutive secretion are also degraded by lysosomes, so crinophagy of vesicles in the constitutive secretory pathway also occurs. Quantification of the amount of a particular secretory protein that is degraded by crinophagy usually requires biochemical analysis using pulse-chase protocols as described earlier (see Chapter 2) with careful accounting of all the newly synthesized protein in question.25 Such studies typically reveal that 20-90% of a newly synthesized secretory protein is degraded rather than secreted (Fig. 3.4). Crinophagy has not been reported in yeast, but it probably exists. Crinophagy has also not been reproduced in any cell free systems.

Constitutive Secretion Pathways Proteins that enter the constitutive secretion pathway are synthesized on polysomes bound to the ER or enter the ER posttranslationally.26 Modifications of the protein including signal peptide cleavage, disulfide bond formation, and glycosylation occur within the ER lumen. Multimeric proteins usually begin their oligomerization within the ER.10,27 The rate of protein movement from the ER to the cis-Golgi is highly variable.27 At least part of the heterogeneity in transport rates is due to interaction of some proteins with molecular chaperones (the glucose-regulated protein of 78 kDa, calnexin, calreticulin, etc.) in the ER lumen.28,29 Other ER lumenal resident proteins contain carboxyl terminal KDELrelated sequences that cause them to be returned to the ER after reaching the cisGolgi or ER-Golgi intermediate compartment (ERGIC).29 This return is mediated by KDEL receptors and vesicles that are coated with co-atomer I proteins (COPI). ER membrane proteins often contain di-basic KK or RR sequences that also bind to COPI. Exit from the ER can also be prevented by inhibitors of ATP production and by temperatures below 15˚C.30 The proteins then travel sequentially to the cis-, medial-, and trans-Golgi stacks and then to the TGN where they may be further glycosylated, sulfated, and/or acylated (Fig. 3.5).31 Proteins travel between Golgi stacks in carrier vesicles coated not with COPI but with co-atomer protein II (COPII) complexes.32,33 Soon after the coated vesicle buds off from the Golgi, the coat is removed to expose v-SNAREs as described in Chapter 2 concerning trafficking of endocytic vesicles. The ARF GTPase is also required for this

44


Fig. 3.4. Representative experimental results of the quantitation of the amount of a protein hormone synthesized, degraded, and secreted. Cells were radiolabeled with [35S]methionine, and the protein hormone was purified using specific antibodies. The radioactivity associated with the hormone was followed. Some of the radiolabel (5%) was lost very rapidly (T1/2 = 25 min) as the protein hormone was processed to its mature size, in this case representing cleavage of the ER signal sequence. The radioactivity in the secreted protein hormone could account for 60% of the hormone that was made. No radioactive hormone could be found within the cell at the end of the experiment, therefore 35% of the synthesized protein hormone must have been degraded.

movement.32,33 The vectorial transport through the cis, medial, and trans-Golgi stacks and the TGN requires the progressive acidification of these compartments.34 It is possible that cargo proteins may have to undergo structural modifications induced by the lower pH before they can move to the next stack. Alternatively, the same v-SNARE may be used in the vesicular trafficking between cis-, medial-, and trans-Golgi, but the v-SNARE may be progressively modified in structure due to the progressive acidification. There are presumably different t-SNAREs in the different Golgi subcompartments. The movement of these secretory vesicles to the plasma membrane requires cytoskeletal elements including actin and tubulin.1,3 There is only a modest (at most 2-fold) concentration of cargo proteins in these vesicles compared to their concentration in the ER lumen.34,35 As mentioned earlier, this pathway appears to be the default pathway. For example, lysosomal proteins that contain mannose-6-phosphate modifications are efficiently sorted to lysosomes by the mannose-6-phosphate receptor at the TGN. If this sorting is blocked, the lysosomal enzymes are secreted from the cell by the constitutive secretion pathway.36 Furthermore, two small molecules that are unlikely to contain targeting information, a glycosaminoglycan and a glycosylated acyltripeptide, are secreted by the constitutive pathway.37,38 The vesicles that bud from the trans-Golgi or TGN for this constitutive secretion pathway contain cargo proteins to be secreted and also membrane-spanning proteins destined for residence in the plasma membrane.39


45

Fig. 3.5. Pathways of protein secretion through the ER and Golgi compartments. Proteins destined for both constitutive and regulated secretion pathways travel from the ER through the ERGIC and then to the cis-, medial-, and trans-Golgi stacks. They are sorted into different vesicles at the TGN. The vesicles surrounding the Golgi which are coated with COPII (O) are responsible for this anterograde movement. Those coated with COPI (O) are responsible for retrograde transport of proteins to the ERGIC and the ER.

Regulated Secretion Pathways Cells that utilize a regulated secretory pathway are easily identified by their abundant, electrondense, secretory granules (Fig. 3.6).7,13 Such cells include endocrine and exocrine cells, neurons, platelets, mast cells, and others. In response to a stimulus that causes an elevation of intracellular calcium or other second messengers, the secretory granules fuse with the plasma membrane, and their contents are released outside the cell. The route traveled by proteins in the regulated secretory pathway appears identical to those in the constitutive secretion pathway until reaching the TGN.1-3 Proteins destined for the regulated secretory pathway are selectively transferred into budding regions of TGN that are coated with clathrin and adaptor protein-1 complexes (AP-1).40 Immature secretory vesicles (ISVs) continue to have clathrin associated with them while the mature secretory vesicles do not. This maturation is accompanied by proteolytic processing of certain secreted proteins and by concentration of the secreted protein by as much as 200-fold above levels found in the ER.1-3 As mentioned earlier, proteins in the constitutive secretory pathway are seldom concentrated more than 2-fold above levels in the ER.1,35 This sorting of proteins into the regulated secretory pathway appears to require specific receptors for such proteins in the TGN.41-43 The targeting signal in the protein is not a particular carbohydrate as is the case for targeting of lysosomal proteins by the mannose6-phosphate receptor, nor a short linear amino acid sequence such as KDEL for ER retention. Instead, the targeting motif appears to be conformation-dependent.12,44 This sorting signal was first identified in pro-opiomelanocortin (POMC), the prohormone for adrenocorticotropin, β endorphin, and α melanocyte-stimulating hormone, synthesized in pituitary cells. Amino acids 8-20 of POMC form an amphipathic loop stabilized by a

46


Fig. 3.6. Two cells that are active in the regulated secretory pathway. The hallmark of such cells are the numerous, dense secretory vesicles (SV). N = nucleus.

disulfide bridge (Fig. 3.7).12,43 The POMC 1-26 peptide was synthesized and radiolabeled with 125I. This ligand specifically bound to a protein associated with TGN membranes.43 The sorting receptor was sequenced and identified as a previously characterized 55 kDa protein, carboxypeptidase E.43 The membrane-associated form of the enzyme has little carboxypeptidase activity, but it is able to bind to POMC, proinsulin, proenkephalin, and prochromogranin-A. This binding is optimal at acidic pH, and both the TGN and ISVs are acidified. When the carboxypeptidase E gene is knocked out in mice, POMC is secreted from pituitary cells by the constitutive pathway but not by the regulated pathway.43 Other membrane proteins have also been proposed to be receptors for sorting proteins into the regulated secretory pathway. Solubilized Golgi membrane proteins from canine pancreas were passed through an affinity column containing immobilized protein precursors that entered the regulated secretory pathway.41 Two or three proteins in the 25 kDa range were recovered. These proteins also bound to insulin and growth hormone, but not to bovine serum albumin, hemoglobin, or myoglobin.41 No direct evidence that these 25 kDa proteins are sorting receptors for the regulated secretory pathway have been reported. An additional consideration about protein targeting to the regulated secretory pathway is that such proteins tend to aggregate in the acidic pH of the TGN and ISV.1-3,46 This aggregation is evident as the intense immunogold labeling of the dense core of the secretory vesicles. This aggregation or condensation may prevent the protein from leaving the ISV. The binding to sorting receptors together with the tendency to aggregate may combine to insure efficient targeting and concentration of proteins in the regulated secretory pathway.47 The fact that protein sorting continues to occur in ISVs has been convincingly demonstrated for pancreatic β cells that secrete insulin.48 In these cells a substantial proportion of newly synthesized lysosomal cathepsins are initially mistargeted to the regulated secretory pathway rather than to lysosomes. Carrier vesicles can bud off the ISVs utilizing AP-1 and clathrin, and these carrier vesicles contain the mannose-6-phosphate receptor and mannose-6-phosphate-


47

Fig. 3.7. The recognition element in POMC that is bound by receptors in the TGN. This peptide region is near the amino terminus of POMC, and the disulfide bond is a required feature for its recognition. Amino acids shown are A, alanine; C, cysteine; D, aspartate; E, glutamate; L, leucine; N, asparagine; Q, glutamine; S, serine; and T, threonine.

marked lysosomal proteins (Fig. 3.8). These vesicles then fuse with early endosomes and/or late endosomes after which the cargo protein is delivered to lysosomes. Such sorting stops by the time the secretory vesicle has reached maturity.48 The proteolytic processing of secretory proteins to their active, mature forms occurs in the TGN or in the ISV. In the case of proinsulin cleavage to insulin, most of the processing is in the ISV.49 As expected the ISV also contains the trypsin-like protease responsible for converting proinsulin to insulin. Interestingly, under conditions where the stimulation for insulin secretion is low, there is a selective secretion of the proinsulin connecting peptide.50 The probable explanation for this finding is that the connecting peptide is shuttled to the constitutive secretory pathway through vesicles derived from the ISV while the mature insulin is retained in the regulated secretory pathway which is inactive when the demand for insulin is low. A substantial amount of new membrane can be delivered to the plasma membrane through the regulated secretory pathway.7 For example, when the exocrine pancreas receives a signal to secrete hydrolases, the plasma membrane area (30 µm2) has been calculated to be theoretically greatly increased (900 µm2). However, this extra membrane is rapidly returned to the trans-Golgi or TGN by endocytosis, so the increase in plasma membrane surface area is barely detectable. In this fashion the membrane components for the regulated secretory pathway may be recycled many times (Fig. 3.9).

48


Fig. 3.8. Mistargeted lysosomal enzymes can be retrieved from immature secretory vesicles (ISV) and delivered to lysosomes through the endocytic pathway. An ISV can form buds which contain AP-1 and clathrin and concentrate mannose-6-phosphate receptors and lysosomal enzyme precursors that have been mistargeted to the regulated secretion pathway. The clathrin-coated vesicles are uncoated by hsc73, and these vesicles fuse with early endosomes and then travel to late endosomes and finally lysosomes (L). PM = plasma membrane; CYT, cytosol.

Fig. 3.9. Fusion of regulated secretory vesicles with the plasma membrane is closely coupled to endocytosis of an equivalent amount of membrane. PM = plasma membrane; CYT = cytosol. Arrows indicate directions of movement of the membrane proteins.

Neurotransmission and Synaptic Vesicle Dynamics The dynamic interaction between endocytosis and exocytosis is especially evident in neurons (Fig. 3.10). In these cells membrane proteins are initially made by the usual route involving transit through the ER and Golgi. Neurotransmitters such as acetylcholine and glutamate are selectively transported into small vesicles called synaptic vesicles after they are formed by the Golgi.51 This transport is driven by a proton gradient due to the presence of a +H-ATPase in the synaptic vesicle membrane.52 A typical axon will contain several hundred synaptic vesicles, but only 20-40 will release their contents into the synapse in


49

Fig. 3.10. Synapses require specialized forms of regulated vesicle secretion. Synaptic vesicles in the axon contain specific transporters for neurotransmitter uptake. The synaptic vesicles are acidic (+), and transport of many neurotransmitters (open arrows) is dependent on proton antiport. The synaptic vesicles partially fuse with the axon plasma membrane to allow the neurotransmitter to enter the extracellular space close to a dendrite. The synaptic vesicle becomes an early endosome even without full mixing of the synaptic vesicle membrane with the plasma membrane. Some of the neurotransmitter may be takenup during endocytosis.

response to a single stimulus.53 These 20-40 synaptic vesicles are docked at specialized plasma membrane domains called active sites. The other synaptic vesicles are associated with cytoskeleton further away from the plasma membrane. Some of these vesicles are released from the cytoskeleton following an action potential and can then be docked at active sites at the plasma membrane.54 The membrane components of synaptic vesicles include the v-SNARE, synaptobrevin, and a protein that is thought to control membrane fusion, synaptotagmin. 55 Synaptotagmin binds calcium and undergoes a conformational change as a result.56 Synaptotagmin may also couple exocytosis and endocytosis since it can bind to AP-2 complexes.56 The production of synaptic vesicles in permeabilized cells requires ATP, cytosol, and GTP hydrolysis.

50


Fig. 3.11. Regulation of crinophagy. N = nucleus; RER = rough endoplasmic reticulum; G = Golgi; SV = secretory vesicle; L = lysosome. A. Pancreatic islet β-cells secreting insulin degrade less insulin by crinophagy when blood glucose concentrations are high. B. In many cell types, inhibition of lysosomal function with weak bases such as ammonium chloride or with inhibitors of lysosomal proteases, such as leupeptin, result in an increase in the amount of protein secreted. C. Colchicine disrupts microtubules and strongly inhibits protein secretion. The amount of the secreted protein that is degraded by crinophagy increases under these conditions.

The synaptic vesicles are triggered to fuse with the plasma membrane by the arrival of an action potential. This action potential opens voltage-gated calcium channels in the axon’s plasma membrane. This region of the plasma membrane also contains t-SNAREs such as syntaxin and SNAP-25. Once again, the v-SNAREs and t-SNAREs not only interact with each other but also recruit NSF and SNAPs required for vesicular fusion. The synaptic


51

Fig. 3.12. Regulated secretory vesicles can also be degraded by macroautophagy. Cells that exhibit regulated secretion show images such as in A, where membrane fusion occurs between a regulated secretory vesicle (SV) and a lysosome (L). The number of such structures increase when the proteins contained in the regulated secretory vesicle are not needed outside the cell. Images such as B are less common and suggest that regulated secretory vesicles can be engulfed in macroautophagic vacuoles (AV) which subsequently fuse with lysosomes to degrade the entire SV.

vesicles may release neurotransmitters without their membranes fully mixing with the plasma membrane (Fig. 3.9). Instead, fusion of a small region of the synaptic vesicle and the plasma membrane may allow the release of neurotransmitters as well as the very efficient recycling of synaptic vesicle membranes. The membrane components of the synaptic vesicles, including transporters for the neurotransmitter, are rapidly retrieved by endocytosis. New synaptic vesicles can form directly from these early endosomes.57

Crinophagy The fusion of both constitutive secretion vesicles and regulated secretion vesicles with lysosomes rather than the plasma membrane results in degradation of the usually secreted proteins (Fig. 3.3). The fraction of the newly synthesized secreted protein that is degraded depends on the protein, the cell type, and the conditions of study. For example, parathyroid hormone acts on bone and kidney to raise plasma calcium levels. Parathyroid hormone

52


Fig. 3.13. Secretory proteins that form granules within the ER lumen can cause the transition of ER to lysosomes. The cell shown is making thyroid stimulating hormone (TSH), the fl-chain of which is within intercisternal granules (ICGs) within the ER lumen. See the text for description of this process. N = nucleus; RER = rough endoplasmic reticulum; ER = endoplasmic reticulum; L = lysosome; PM = plasma membrane.

secretion is activated when the serum calcium concentration falls below 10 mM. The fraction of parathyroid hormone degraded by crinophagy ranges from 40% when serum calcium levels are low to 90% when calcium concentrations are above 10 mM.24,58 Other endocrine tissues also exhibit some degree of crinophagy. Anterior pituitary cells that synthesize and secrete prolactin rapidly degrade prolactin-containing secretory granules in response to cessation of breast feeding.4,59 When insulin-secreting β cells are in the presence of high glucose concentrations, most of the insulin is secreted. However, when glucose concentrations fall, 25% of the insulin-containing secretory granules fuse


53

with lysosomes (Fig. 3.11).60 Similarly, when insulin secretion was blocked with diazoxide, crinophagy of β cell granules was induced (Fig. 3.11).61 Other cell types also degrade a significant fraction of their secretory proteins. Mammary glands synthesize and secrete casein, the major protein in milk, but whether this secretion is by the regulated or the constitutive pathway remains unclear. Most (90%) of the newly synthesized casein in mammary gland organ culture is degraded rather than secreted.62 Whether or not this large percentage of degradation also applies in mammary glands within intact animals remains to be determined. Collagen is secreted by fibroblasts by the constitutive pathway, and 15% of the newly synthesized collagen is degraded by crinophagy.63 Hepatocytes constitutively secrete a variety of serum proteins, but 40% of the newly synthesized proteins are degraded in lysosomes by crinophagy.64 Although some variability was found for the different secreted proteins analyzed, this result applied to serum albumin, transferrin, and α2-microglobulin. As expected, much of the degradation of secreted protein by crinophagy can be inhibited by lysosomotropic agents such as NH4Cl, chloroquine, and primacrine.64,65 Degradation by crinophagy can also be partially inhibited by cysteine protease inhibitors such as leupeptin. In response to these inhibitors, the amount of protein secreted increases (Fig. 3.11).64,65 In addition, microtubule inhibitors like colchicine and vinblastine preferentially inhibit fusion of secretory granules with the plasma membrane (Fig. 3.11). In response to these inhibitors, crinophagy increases.65 However, microtubule inhibitors also stimulate macroautophagy,66 so the interpretation of results after disruption of microtubules is problematic (see below). Forms of crinophagy in addition to that described so far have also been reported (Fig. 3.12). For example, in rat liver entire secretory vesicles can be engulfed by macroautophagic vacuoles prior to fusion with lysosomes.65 The major difference in the result of this process compared to the direct fusion of secretory vesicles with lysosomes is that membrane components of the vesicles as well as cargo should be delivered to lysosomes and degraded. The direct fusion of secretory vesicles with lysosomes leaves open the possibility of resequestering and recycling of the membrane components. Another form of crinophagy has been reported in anterior pituitary cells that make thyroid stimulating hormone (TSH).59 When animals are thyroidectomized to relieve the negative feedback inhibition of TSH production by thyroid hormones, TSH synthesis markedly increases. This increase in synthesis is accompanied by swelling of the ER lumen and the production of intercisternal granules (ICGs) that contain the β subunit of TSH. When TSH production is inhibited by the injection of thyroid hormones, the ICG are degraded by a mechanism that involves transfer of ICGs to transitional regions between rough and smooth ER, thickening of the surrounding membrane, acquisition of lysosomal membrane proteins, acquisition of lysosomal matrix proteins, degradation of the ICG and TSH β chains, and production of a lysosome (Fig. 3.13).59 The extent to which this pathway operates under normal conditions and in other cell types is not known. It seems possible that this pathway is activated only when ICGs form.

Future Directions of Research The identification of receptors in addition to carboxypeptidase E for packaging of proteins into regulated secretory vesicles requires further study. The subcellular distribution of such receptors as well as regulation of their activity by posttranslational modifications and interactions with other proteins require further study. The receptor half-lives and site

54


of degradation will also increase our understanding of regulated protein secretion and crinophagy. Fusion of vesicles in the constitutive secretion pathway with the vacuole rather than the plasma membrane should be explored in yeast. If crinophagy exists in yeast, genetic screens may be designed to identify mutations in the crinophagic process. Factors that actually regulate the fusion of secretory granules with lysosomes rather than the plasma membrane are not known. However, it is reasonable to speculate that t-SNAREs and v-SNAREs may be differentially modified depending on the levels of second messengers used to trigger secretory granule fusion with the plasma membrane. For example, the t-SNARE on the lysosome may be blocked, or otherwise made inactive, under conditions in which secretion of the protein is activated. In the absence of secretory signals, the t-SNARE on the lysosome membrane may be activated while that in the plasma membrane may be inactivated. Molecular details about crinophagy will undoubtedly require the reconstitution of crinophagy with purified lysosomes and secretory granules. With such a system requirement for cytosol, ATP, and other co-factors can be determined. In addition, the interacting v-SNARE on the secretory granule and the t-SNARE on the lysosome may be identified. The processes by which ICGs are degraded by the transition of ER into lysosomes also leaves several important areas to be studied further.59 For example, how the ER membrane receives lysosomal membrane and matrix constituents is unclear. If vesicles derived from the TGN that normally traffic proteins to lysosomes are diverted to fuse with ER, the machinery regulating this change in destination requires study References 1. Burgess L, Kelly RB. Constitutive and regulated secretion of proteins. Ann Rev Cell Biol 1987; 3:243-293. 2. Pryer NK, Wuesthube LJ, Schekman R. Vesicle-mediated protein sorting. Ann Rev Biochem 1992; 61:471-516. 3. Burgouyne RD, Morgan A. Regulated exocytosis. Biochem J 1993; 293:305-316. 4. Farquhar MG. Secretion and crinophagy in prolactin cells. Adv Exp Med Biol 1977; 80:37-94. 5. Glaumann H. Crinophagy as a means for degrading excess secretory proteins in rat liver. Revis Biol Celular 1989; 20:97-110. 6. Kuriakose NR, Reifel CW, Bendayan M et al. Prolactin crinophagy is induced in the estrogenstimulated male rat pituitary. Histochem 1989; 92:499-503. 7. Palade G. Intracellular aspects of the process of protein synthesis. Science 1975; 189:347-358. 8. Salpeter MM, Farquhar MG. High resolution analysis of the secretory pathway in mammotrophs of the rat anterior pituitary. J Cell Biol 1981; 91:240-246. 9. Griffiths G, Pfeiffer S, Simons K et al. Exit of newly synthesized membrane proteins from the trans cisterna of the Golgi complex to the plasma membrane. J Cell Biol 1985; 101:949-964. 10. Kelly RB. Pathways of protein secretion in eukaryotes. Science 1985; 230:25-32. 11. Stevens TH, Rothman JH, Payne G et al. Gene dosage-dependent secretion of yeast vacuolar carboxypeptidase Y. J Cell Biol 1986; 102:1551-1557. 12. Cool DR, Loh YP. Identification of a sorting signal for the regulated secretory pathway at the N-terminus of pro-opiomelanocortim. Biochimie 1994; 76:265-270. 13. Valls LA, Hunter CP, Rothman JH et al. Protein sorting in yeast. Localization determinant of yeast vacuolar carboxypeptidase Y resides in the propeptide. Cell 1987; 48:887-897. 14. Moore H-P H, Walker MD, Lee F et al. Expressing a human proinsulin cDNA in a mouse ACTHsecreting cell. Intracellular storage, proteolytic processing, and secretion on stimulation. Cell 1983; 35:531-538. 15. Comb M, Liston D, Martin M et al. Expression of the human proenkephalin gene in mouse pituitary cells. Accurate and efficient mRNA production and proteolytic processing. EMBO J 1985; 4:3115-3122. 16. Martin TF, Walent H. A new method for cell permeabilization reveals a cytosolic requirement for Ca+2-activated secretion in GH3 pituitary cells. J Biol Chem 1989; 264:10299-10308.


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17. Grimes M, Kelly RB. Intermediates in the constitutive and regulated secretory pathway released in vitro from semi-intact cells. J Cell Biol 1992; 117:539-549. 18. Desnos C, Clift-O’Grady L, Kelly RB. Biogenesis of synaptic vesicles in vitro. J Cell Biol 1995; 130:1041-1049. 19. Creutz CE. The annexins and exocytosis. Science 1992; 258:924-931. 20. Novick P, Schekman R. Secretion and cell-surface growth are blocked in a temperature-sensitive mutant of Saccharomyces cerevisiae. Proc Nat Acad Sci USA 1979; 76:1858-1862. 21. Novick P, Field C, Schekman R. Identification of 23 complementation groups required for posttranslation events in the yeast secretory pathway. Cell 1980; 21:205-215. 22. Bennett MK, Scheller RH. The molecular machinery for secretion is conserved from yeast to neurons. Proc Nat Acad Sci USA 1993; 90:2559-2563. 23. Chertow BS. The role of lysosomes and proteases in hormone secretion and degradation. Endocr Res 1981; 2:137-173. 24. Morrissey JJ, Cohn DV. Secretion and degradation of parathormone as a function of intracellular maturation of hormone pools. J Cell Biol 1979; 83:521-528. 25. Yokota S, Fahimi HD. Immunocytochemical localization of albumin in the secretory apparatus of rat liver parenchymal cells. Proc Nat Acad Sci USA 1981; 78:4970-4974. 26. Schatz G, Dobberstein B. Common principles of protein translocation across membranes. Science 1996; 271:1519-1525. 27. Parent JB, Bauer HC, Olden K. Three secretory rates in human hepatoma cells. Biochim Biophys Acta 1985; 846:44-50. 28. Hauri H-P, Schweitzer A. The endoplasmic reticulum-Golgi intermediate compartment. Curr Opin Cell Biol 1992; 4:600-608. 29. Pelham HR. Recycling of proteins between the endoplasmic reticulum and Golgi complex. Curr Opin Cell Biol 1991; 3:585-591. 30. Tartakoff AM. Temperature and energy depedence of secretory protein transport in the exocrine pancreas. EMBO J 1986; 5:1477-1482. 31. Balch WE. Molecular dissection of early stages of the eukaryotic secretory pathway. Curr Opin Cell Biol 1990; 2:634-641. 32. Kreis TE. Regulation of vesicular and tubular membrane traffic of the Golgi complex by coat proteins. Curr Opin Cell Biol 1992; 4:609-615. 33. Pepperkok R, Scheel J, Horstmann H et al. Beta COP is required for biosynthetic membrane transport in the endoplasmic reticulum to the Golgi complex in vivo. Cell 1993; 74:71-82. 34. Mellman I, Fuchs R, Helenius A. Acidification of the endocytic and exocytic pathways. Annu Rev Biochem 1986; 55:663-700. 35. Hearn SA, Silver MM, Sholdice JA. Immunoelectron microscopic labeling of immunoglobulins in plasma cells after osmium fixation and epoxy embedding. J Histochem Cytochem 1985; 33:1212-1218. 36. Dong J, Prence EM, Sahagian GG. Mechanism for selective secretion of a lysosomal protein by transformed mouse fibroblasts. J Biol Chem 1989; 264:7377-7383. 37. Burgess TL, Kelly RB. Sorting and secretion of adrenocorticotropin in a pituitary tumor cell line after perturbation of the level of a secretory-specific proteoglycan. J Cell Biol 1984; 99:2223-2230. 38. Wieland FT, Gleason MI, Serafini T et al. The rate of bulk flow from the endoplasmic reticulum to the cell surface. Cell 1987; 50:289-300. 39. Holcomb CL, Hansen WB, Eicheverry TE et al. Plasma membrane protein intermediates are present in the secretory vesicles of yeast. J Cell Biol 1988; 106:641-648. 40. Bruzzone R. The molecular basis of enzyme secretion. Gastroenterology 1990; 99:1157-1176. 41. Chung K-N, Walter P, Aponte G et al. Molecular sorting in the secretory pathway. Science 1989; 243:192-197. 42. Calakos N, Bennett MK, Peterson KE et al. Protein-protein interactions contribute to the specificity of intracellular vesicle trafficking. Science 1994; 263:1146-1149. 43. Cool DR, Normant E, Shen F-S et al. Carboxypeptidase E is a regulated secretory pathway sorting receptor: Genetic obliteration leads to endocrine disorders in Cpefat mice. Cell 1997; 88:73-83. 44. Chanat E, Weiss U, Huttner WB et al. Reduction of the disulfide bond of chromogranin β (secretogranin 1) in the trans Golgi network causes its missorting to the constitutive secretory pathway. EMBO J 1993; 12:2159-2168. 45. Tam WHH, Andreasson KA, Loh YP. The amino-terminal sequence of pro-opiomelanocortin directs intracellular targeting to the regulated secretory pathway. Eur J Cell Biol 1993; 62:294-306.

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46. Orci L, Ravazzola M, Amherdt M et al. The trans-most cisterna of the Golgi complex: A compartment for sorting of secretory proteins and plasma membrane proteins. Cell 1987; 51:1039-1051. 47. Farquhar MG, Palade GE. The Golgi apparatus: 100 years of progress and controversy. Trends Cell Biol 1998; 8:2-10. 48. Kuliawat R, Klumperman J, Ludwig T et al. Differential sorting of lysosomal enzymes out of the regulated secretory pathway in pancreatic β-cells. J Cell Biol 1997; 137:595-608. 49. Orci L, Ravazzola M, Amberdt M et al. Direct identification of prohormone conversion site in insulin-secreting cells. Cell 1985; 42:671-681. 50. Rhodes CJ, Halban PA. Newly synthesized proinsulin-insulin as well as stored insulin are released from pancreatic β-cells uniquely via a regulated, not a constitutive, pathway. J Cell Biol 1987; 105:145-153. 51. Jahn R, Südhof TC. Synaptic vesicles and exocytosis. Annu Rev Neurosci 1994; 17:219-246. 52. Forgac M. Regulation of vacuolar acidification. Soc Gen Physiol Ser 1996; 51:121-132. 53. McClure SJ, Robinson PJ. Dynamin, endocytosis, and intracellular signalling. Mol Memb Biol 1996; 13:189-215. 54. Trifaro J-M, Vitale ML. Cytoskeleton dynamics during neurotransmitter release. Trends Neurosci 1993; 16:466-472. 55. Littleton JT, Bellen HJ. Synaptotagmin controls and modulates synaptic vesicle fusion in a Ca+2dependent manner. Trends Neurosci 1995; 18:177-183. 56. Zhang JZ, Davletov BA, Südhof TC et al. Synaptotagmin is a high affinity receptor for clathrin AP-2: Implications for membrane recycling. Cell 1994; 78:751-760. 57. Ryan TA, Smith SJ. Vesicle pool mobilization during action potential firing at hippocampal synapses. Neuron 1995; 14:983-989. 58. Habener JF, Kemper BW, Rich A et al. Biosynthesis of parathyroid hormone. Rec Prog Horm Res 1977; 33:249-308. 59. Noda T, Farquhar MG. A non-autophagic pathway for diversion of ER secretory proteins to lysosomes. J Cell Biol 1992; 119:85-97. 60. Schnell AH, Swenne I, Borg LA. A quantitative estimation of crinophagy in the mouse pancreatic β-cell. Cell Tissue Res 1988; 252:9-15. 61. Skoglund G, Ahren B, Lundquist I. Biochemical determination of islet cell lysosomal enzyme activities following crinophagy-stimulating treatment with diazoxide in mice. Diabetes Res 1987; 6:81-84. 62. Razooki-Hasan H, White DA, Mayer FJ. Extensive destruction of newly synthesized casein in mammary explants in organ culture. Biochem J 1982; 202:133-138. 63. Bienkowski RS. Intracellular degradation of newly synthesized secretory proteins. Biochem J 1983: 214:1-10. 64. Lenk SE, Fisher DL, and Dunn WA. Regulation of protein secretion by crinophagy in perfused rat liver. Eur J Cell Biol 1991; 56:201-209. 65. Marzella L, Glaumann H. Autophagy microautophagy and crinophagy as mechanisms for protein degradation. In: Glaumann H, Ballard FJ, eds. Lysosomes: Their Role in Protein Breakdown. New York: Academic Press, 1987:319-367. 66. Grinde B and Seglen PO. Role of microtubuli in the lysosomal degradation of endogenous and exogenous protein in isolated rat hepatocytes. Hoppe-Seylerís Z Physiol Chem 1981; 362:549-556.

CHAPTER 4

Degradation of Intracellular Proteins by Macroautophagy Overview

M

any cells are able to degrade intracellular proteins by macroautophagy.1-3 In this multistep process, regions of cytoplasm are surrounded by a double membrane to form an organelle called an early autophagosome.1,4 The material sequestered in such autophagosomes includes recognizable organelles such as mitochondria, peroxisomes, ribosomes, and glycogen granules, as well as a variety of cytosolic proteins in proportion to their abundance in the cytosol (Table 4.1).5,6 Large cytoskeletal elements as well as nuclear structures are excluded from autophagosomes.1-3 The origin of the two membranes that surround the autophagosome remains uncertain despite extensive studies over the past 30 years.1-3,7-9 The autophagosome then acquires the membrane ATP-dependent proton pump, as well as certain other lysosomal membrane proteins within its outer-membrane, and becomes acidic.1-3 These structures still contain two membranes and are called late autophagosomes.1,4 The early and late autophagosomes together are also referred to as autophagic vacuoles. The late autophagosomes then acquire lysosomal hydrolases and additional lysosomal membrane proteins by fusion with primary or secondary lysosomes.1-3 Simultaneously, the innermembrane of the late autophagosome is digested to form a structure called an autolysosome.1,4 The autolysosome is more acidified than the late autophagosome, and in the autolysosome the contents of the autophagic vacuole are digested (Fig. 4.1). Each of these steps in macroautophagy requires ATP.1-3 Rates of macroautophagy in liver are regulated by amino acids and hormones.1-3 In the perfused rat liver leucine is an especially potent regulator of macroautophagy, but seven additional amino acids are required for maximal suppression of macroautophagy.2,10,11 Insulin inhibits while glucagon stimulates macroautophagy.1,3,10,11 These considerations explain why macroautophagy is stimulated in liver between meals when circulating amino acid levels fall and the insulin/glucagon ratio is low. Regulation of macroautophagy in other tissues is somewhat different from liver. Some cells in culture stimulate autophagy in the absence of amino acids,12 similar to the situation with liver. However, fibroblasts in culture increase macroautophagy when they reach confluence13-15 even in the presence of amino acids. Intracellular regulators of macroautophagy are not yet completely known but already appear to involve multiple second messenger pathways. Intracellular calcium is required for macroautophagy, and a role for calmodulin has been proposed.16 Phosphorylation of a


58


Fig. 4.1. The multistep process of macroautophagy. (A) The double membrane surrounding the autophagic vacuole has been proposed to be derived from rough ER (RER), smooth ER (SER), the Golgi (G), or an unique organelle, the phagophore (P). (B) The autophagic vacuole (AV) digests its inner membrane, and the proton pumping ATPase makes the AV acidic. + = proton. (C) The AV acquire lysosomal hydrolases by fusion with primary or secondary lysosomes (L). M = mitochondrion.

cytokeratin of 52 kilodaltons (kDa) may inhibit macroautophagy,17 and this phosphorylation is in turn controlled by a kinase and a phosphatase. Other phosphorylated proteins, such as the S6 ribosomal protein,18 may inhibit macroautophagy. There is also growing evidence that heterotrimeric G proteins stimulate macroautophagy in undifferentiated colon cancer cells.19 Finally, inhibitors of phosphotidylinositol-3-kinase (PI3 kinase) inhibit macroautophagy in hepatocytes.20

Degradation of Intracellular Proteins by Macroautophagy

59

Fig. 4.2. Correlation between the volume of autophagic vacuoles and rates of proteolysis in the perfused rat liver. Rates of proteolysis were measured as the release of [14C]valine from livers previously radiolabeled by intraperitoneal injection of the isotope. Protein degradation is expressed as fractional degradation rate per hr. Both early and late autophagosomes were quantitated by electron microscopy under the variety of conditions examined. These conditions included perfusion in the presence of varying amounts and composition of amino acids and in response to refeeding of starved animals.

Methods Used to Study Macroautophagy Ultrastructural studies at the electron microscope level have been very important in the study of macroautophagy.1-3,4,7,8,14 The variable size and morphology of autophagic vacuoles can make these structures difficult to quantify, and certain autophagic vacuoles without recognizable intracellular structures can be easily missed. Nevertheless, Mortimore and his colleagues2,3,10,11 have extensively examined protein degradation using perfused rat liver and have been able to quantitate numbers and areas of autophagic vacuoles as well as rates of proteolysis under a variety of conditions (Fig. 4.2). This work clearly indicates that macroautophagy accounts for most of the degradation of long-lived proteins in liver after acute withdrawal of amino acids. The use of pharmacological agents in perfused liver and in cultured hepatocytes has implicated roles for the cytoskeleton in various stages of macroautophagy3 and, more recently, roles for GTPases19 and phosphotidylinositols20 in regulating the formation of autophagic vacuoles. These results must be viewed with some caution since the proposed degree of specificity of drugs is usually inversely related to the number of studies using that drug.

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Fig. 4.3. Hypothetical relationship between the proposed selectivity of a drug’s action and the number of studies of that drug. The graph indicates that no drug is completely selective. 84

Furthermore, it is not always clear whether a particular drug is directly or indirectly affecting macroautophagy. One example of a compound that inhibits the formation of autophagosomes is 3-methyladenine.21 However, this compound also affects aspects of endocytosis.22 Recently, 3-methyladenine has been shown to inhibit PI3 kinase, an enzyme required for many stages of vesicular traffic,23 including endocytosis, exocytosis, and targeting of proteins to the lysosome/vacuole. This increase in the sites of action of a drug with increasing numbers of studies is illustrated in Figure 4.3. Another important technique developed by Seglen and co-workers2,24 to study macroautophagy in hepatocytes is to transiently permeabilize cells to allow [14C]sucrose to enter the cytosol. This sugar cannot be transported into intracellular organelles nor can it be metabolized within lysosomes, so its entry into the lysosomal compartment reflects the rate of nonselective macroautophagy. In isolated hepatocytes 6-7% of the cytosolic [14C]sucrose is transferred to lysosomes per hour. This rate of transfer is markedly reduced by amino acids and by 3-methyladenine suggesting that the transfer process is due to macroautophagy (Fig. 4.4). The use of yeast to study macroautophagy utilizes the combined power of biochemical and genetic approaches. The isolation of several different yeast mutants that are defective in macroautophagy25,26 has greatly expanded our understanding of mechanisms and regulation of macroautophagy. Mutants that fail to stimulate macroautophagy in response


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Fig. 4.4. [14C]sucrose captured by lysosomes. Hepatocytes were electropermeabilized in the presence of [14C]sucrose. The rate of capture of the [14C]sucrose by lysosomes was determined by isolating autolysosomes at increasing times after electropermeabilization. The effects of serum levels of 20 amino acids and 3methyladenine (3-MA) are shown.

to nitrogen starvation are called apg. At least 14 different complementation groups have been defined, and representative cDNAs have been sequenced. Most APG genes are novel and are required for macroautophagy and survival under starvation conditions but not for vegetative growth in rich medium. One mutant, apg1, lies in a gene encoding a serine/threonine protein kinase,27 and another, apg-13, is in a gene encoding a hyperphosphorylated protein.28 Most interestingly, these apg mutants partially overlap with mutants in a cytosol-to-vacuole (ctv) targeting pathway for the vacuolar resident protein, aminopeptidase I (see below).29 An impediment to progress in understanding macroautophagy has been that, until recently, no cell-free systems which reproduce this complex process have been developed. Hepatocyes that have been permeabilized with the α toxin of Staphylococcus aureus will form autophagosomes, and, under the right conditions, will mature the autophagosomes to autolysosomes.30 Both the formation and the maturation require cytosol and ATP and are inhibited by GTP-g−S.30

Tissue Specificity of Macroautophagy Macroautophagy has been most thoroughly studied in liver but probably occurs to some extent in all eukaryotic cells.1-3 An increase in number and volume of macroautophagic

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vacuoles under conditions of increased protein degradation has been demonstrated in liver,1-3 kidney,31 heart,32 skeletal muscle,33 and brain,34 as well as in many cell types in culture.13-15 However, most quantitative work has been with liver. The more recent studies of autophagy in yeast underscores the universality of this process.27-29 Macroautophagy accounts for most of intracellular protein degradation in liver between meals,35 but the majority of protein breakdown in lymphocytes and skeletal muscle may be through the ubiquitin/proteasome pathway. 36 Therefore, the importance of macroautophagy in contributing to total protein degradation will depend on the cell type and the conditions analyzed.

Mechanisms of Macroautophagy The double membrane that surrounds autophagic vacuoles has been proposed to be derived from the Golgi,7 the smooth endoplasmic reticulum (SER),8 the rough endoplasmic reticulum (RER),9 or a distinct lipid-rich organelle, the phagophore.2,37 These conclusions were based on both structural and biochemical analyses. For example, membrane continuities between Golgi and autophagic vacuoles were observed by Novikoff and Shin,7 but Dunn8 reported that autophagic vacuole membranes contain protein markers evident in ribosome-free regions of RER, and Ueno and co-workers9 reported that autophagosomes contained proteins typical of SER. Seglen and co-workers2,37 found autophagic membrane proteins to be distinct from proteins comprising any preexisting membrane system, leading them to propose de novo formation of autophagic vacuole membranes or derivation from a distinct organelle, the phagophore. The more recent work does not support a role for intact Golgi in autophagosome formation.38 When Golgi were disrupted by brefeldinA treatment, secretion of proteins was inhibited but autophagic sequestration was not affected. Clearly, use of membranes from the fragmented Golgi to form the autophagosome membrane cannot be ruled out by these experiments.38 Recent work in yeast, Saccharomyces cerevisiae, has investigated protein content of autophagic vacuole membranes using freeze-replica electron microscopic analysis.39 These studies indicate that the inner membrane of autophagic vacuoles is very lipid-rich and almost devoid of proteins. Furthermore, the outer membrane contains only a small amount of protein. These results raise the possibility that the putative membrane proteins immunolocalized to autophagosomes may represent material sequestered within the autophagosome rather than components of its own limiting membrane. This possibility seems to be strengthened by the recent report that phagosome membranes contain marker proteins of RER, SER, Golgi, and endosomes.40 Additional studies will be required to resolve the origin of the autophagosome membrane. The new autophagosome matures into a degradative organelle in a stepwise fashion.1 The outer membrane of the autophagosome acquires certain lysosomal membrane proteins including the multimeric ATP-dependent proton pump presumably by fusion with vesicles deficient in lumenal hydrolytic enzymes. The origin of such vesicles may be the trans Golgi network (TGN), but little is known about how such putatively empty vesicles may be formed. This late autophagosome is further modified by the acquisition of lysosomal hydrolases most likely by fusion with primary or secondary lysosomes.1,8 These vesicular fusion events will undoubtedly be regulated by protein-protein interactions as in other aspects of vesicular traffic within cells. For example, fusion of the two organelles probably requires the NEM-sensitive fusion protein (NSF), soluble NSF attachment proteins


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Fig. 4.5. Possible roles of ubiquitination in macroautophagy. N = nucleus; EA = early autophagosome; LA = late autophagosome; AL = autolysosome; L = lysosome; sub = substrate-ubiquitin conjugate; i-ub, inhibitor of AL formation-ubiquitin conjugate; ub = ubiquitin. 1. Ubiquitin conjugation may be required for some substrates to be targeted to early autophagosomes. 2. Ubiquitin conjugation to LA or L membrane proteins may be required for fusion of AL and L. 3. An inhibitor of AL and L fusion may be a substrate for ubiquitin mediated degradation. If ubiquitin conjugation is impaired, the level of this putative inhibitor would rise.

(SNAPs), and SNAP receptors (SNAREs) in the lysosomal membrane and the autophagosome outer membrane. However, none of these components have yet been identified. Ubiquitin conjugation is required for the maturation of autophagosomes to autolysosomes.41 Ubiquitin and ubiquitin-protein conjugates are abundant within lysosomes,42,43 and cells containing a temperature-sensitive ubiquitin-activating enzyme (E1) failed to increase protein degradation in response to amino acid deprivation.44 Further analysis showed that autophagosomes form but do not mature to autolysosomes when E1 is inactivated.41 The requirement for ubiquitination may be to mark certain substrate proteins prior to their entry into autophagosomes and/or for fusion of late autophagosomes with lysosomes. Perhaps a short-lived protein inhibits autophagosome-lysosome fusion and this protein is normally kept at low concentrations due to degradation by the ubiquitin/ proteasome pathway. These possible roles for ubiquitination in macroautophagy are diagrammed in Figure 4.5. The autophagosomes rapidly form and mature under normal circumstances. When macroautophagy is suppressed by the addition of amino acids, autophagic vacuoles in liver decline with a half-life of eight min (Fig. 4.6). 4,45 Therefore, flux through the macroautophagic pathway may be great even though the fractional volume accounts for no more than 1.5% of a mammalian cell. Macroautophagy in yeast appears to follow similar steps as those described above for mammalian cells. In this case, however, the autophagosomes fuse with the single yeast vacuole rather than with smaller sized lysosomes of mammalian cells.

Regulation of Macroautophagy As mentioned earlier amino acids strongly inhibit macroautophagy in perfused liver3,10,11 and in isolated hepatocytes2 as well as in heart,32 skeletal muscle,33 and certain

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Fig. 4.6. Rate of disappearance of autophagic vacuoles from perfused rat liver. Macroautophagy was abruptly inhibited by inclusion of amino acids in the perfusion medium. The volume of autophagic vacuoles was quantitated by electron microscopy at increasing times. The autophagic vacuoles declined with a half-life of eight min.

other cell types in culture.13-15 Not all amino acids are effective in this inhibition; in liver leucine is the most effective single amino acid, but maximal suppression of macroautophagy also requires phenylalanine, tyrosine, glutamine, proline, histidine, tryptophan, and methionine.3,10 In addition, alanine does not itself affect macroautophagy but is a co-regulator in the presence of the other amino acids.3,10 Maximal inhibition of macroautophagy in kidney can be achieved with leucine and phenylalanine only,31 inhibition in skeletal muscle is due to leucine and glutamine only,32 and inhibition in heart is due to leucine only.33 A comparison of regulatory amino acids that suppress macroautophagy in different tissues is shown in Figure 4.7.


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In Saccharomyces cerevisiae macroautophagy is induced by nutrient deprivation. Starvation for either nitrogen or carbon activates macroautophagy. Other types of nutritional shifts activate macroautophagy in different types of yeast. For example, the methylotrophic yeast, Pichia pastoris, activates macroautophagy in response to changing the carbon source from methanol to ethanol while a shift from methanol to glucose activates microautophagy (see Chapter 5). The initial formation of autophagosomes seems to be the step of macroautophagy that is most inhibited by amino acids.46 This conclusion was particularly well demonstrated by Seglen and co-workers who followed the sequestration of [14C]sucrose into lysosomes following electro-permeabilization of hepatocytes.24,46 However, amino acids also inhibit steps of macroautophagy after initial formation of autophagosomes.47 For this reason the kinetics of loss of autophagic vacuoles after addition of amino acids (Fig. 4.6) may be somewhat slower than their rate of maturation in the absence of amino acids. The mechanism of inhibition of liver protein breakdown by leucine has received considerable attention.3,48-50 Transamination of leucine is very minor in the liver, so leucine metabolic products are not likely to act as regulators.3 The hydroxyl analog of leucine, α-hydroxylisocaproate, acts as well as leucine in inhibiting macroautophagy.48 This result rules out leucyl-tRNA as a regulator of macroautophagy because α-hydroxylisocaproate is not a substrate for leucyl-tRNA synthetases. The intracellular site at which leucine acts to inhibit macroautophagy appears to be at the plasma membrane.49 Circumstantial evidence for this possibility was obtained when isovaleryl-carnitine was shown to be as effective as leucine in inhibiting macroautophagy.49 This leucine analog is rapidly metabolized within hepatocytes so that its intracellular level is only 10% that of leucine. Presumably the concentrations of leucine and isovalerylcarnitine would be equivalent at the plasma membrane just as it is in the extracellular medium. More direct support for a plasma membrane site of action for leucine has come from the synthesis of multiple antigen protein (MAP) with 8 leucines attached to its N termini (Leu8-MAP).50 Leu8-MAP is not metabolized and does not enter the cell, but it is as effective as leucine in inhibiting macroautophagy. Further studies with an iodinated azide derivative of Leu8-MAP have identified a 340 kDa plasma membrane protein that interacts with leucine, but not with isoleucine.51 Unfortunately, the ability of Leu8-MAP to inhibit macroautophagy in hepatocytes could not be reproduced by another research group,52 so the reasons for these discrepancies will have to be worked out before the significance of these results can be assessed. Another finding that may be related to amino acid regulation of macroautophagy is that cycloheximide and other protein synthesis inhibitors strongly suppress macroautophagy in mammalian cells.3,52 This result may be due to amino acids becoming more readily available when protein synthesis is inhibited. Alternatively, the macroautophagic processes may require the continued synthesis of short-lived proteins. Macroautophagy in yeast is not inhibited by cycloheximide, so a difference in the mechanisms or regulation of macroautophagy exists between mammalian cells and yeast. Glucagon increases and insulin inhibits macroautophagy at physiological amino acid concentrations.3,54-55 At high amino acid concentrations glucagon has no effect on macroautophagy.3,55 These results suggest that glucagon and low amino acids stimulate macroautophagy by mechanisms that share at least one critical component. One possibility is that glucagon treatment stimulates amino acid efflux from the liver to actually create an intracellular amino acid deprivation.55 A second possibility is that occupied glucagon

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Fig. 4.7. Amino acids that inhibit macroautophagy in different organs. The amino acids are listed using their single letter notations. L, leucine; Q, glutamine; F, phenylalanine; Y, tyrosine; P, proline; W, tryptophan; H, histidine; M, methionine; A, alanine. A black box indicates that that particular amino acid contributes to the inhibition of macroautophagy in the indicated organ.

receptors in the plasma membrane and unoccupied amino acid receptors in the plasma membrane produce the same second messengers responsible for the stimulation of macroautophagy. One of the second messengers that is elevated due to glucagon interacting with its receptor in the hepatocyte plasma membrane in cyclic AMP (cAMP).56 cAMP by itself will stimulate macroautophagy in the presence of physiological concentrations of amino acids.57 Insulin inhibits macroautophagy even in the presence of excess amino acids.10,53 This result suggest that insulin acts, at least partially, through mechanisms not shared with amino acids. Whether or not the tyrosine kinase activity of the insulin receptor is necessary for this effect on macroautophagy remains to be established. PI3 kinase is also activated in response to insulin binding to its receptor, and this kinase has been shown to inhibit macroautophagy.20 Finally, it is also possible that insulin affects microautophagy (see Chapter 5) or the direct transport of proteins into lysosomes (see Chapter 6) rather than macroautophagy. The cytoskeleton appears to be required for different stages of macroautophagy (Fig. 4.8). For example, cytochalasin B, which disrupts microfilaments, prevents the formation of early autophagosomes (Fig. 4.8).58 Microtubule disrupters like colchicine and vinblastine do not affect the formation of autophagosomes but inhibit the fusion of autophagosomes and lysosomes.59 Connections between lysosomes and microtubules have been described,60 and it is possible that lysosomes and autophagosomes are brought together on microtubule tracks. However, it is also possible that cytoskeletal disrupters inhibit steps of macroautophagy due to secondary effects on cell metabolism and/or membrane transporters. An increase in cell volume or cell swelling inhibits macroautophagy and has a general anabolic effect on liver cell metabolism.61,62 The mechanism(s) of this inhibition are not known, but certain amino acids enter liver cells by transporters that co-transport Na+, and this transport may be associated with increased cell swelling.63 Perhaps these amino acids inhibit macroautophagy, at least in part, due to their effect on cell swelling. Alterations in cell swelling or volume may also cause changes in the cytoskeleton that result in an inhibition of macroautophagy. Finally, increased cell swelling is also associated with changes in


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Fig. 4.8. Effects of inhibitors on macroautophagy. EA = early autophagosome; LA = late autophagosome; L = lysosome; AL = autolysosome; N = nucleus; + = proton. Amino acids and the microfilament disrupter, cytochalasin B, block formation of early autophagosomes, while microtubule disrupters, cholchicine and vinblastine, block the fusion of late autophagosomes with lysosomes. thereby preventing the formation of autolysosomes.

intracellular ion concentrations. In hepatocytes cell swelling is associated with increased intracellular Ca+2. This consideration cannot explain the reduction in macroautophagy, however, because elevated Ca+2 stimulates macroautophagy.16 Changes in other intracellular ions that modify rates of macroautophagy remain to be discovered. The stimulation of macroautophagy by cAMP and Ca+2 suggests that protein phosphorylation events may regulate macroautophagy.64 This conclusion is further supported by findings that several inhibitors of tyrosine protein kinases inhibit macroautophagy.16,65 However, other protein phosphorylation events may inhibit autophagy. For example, inhibitors of Ca+2/calmodulin-dependent protein kinase II stimulate autophagy.66 There is also evidence that phosphorylation of ribosomal protein S6 is correlated with an inhibition of macroautophagy.16,67 The inverse correlation between protein degradation rates and S6 phosphorylation over varying amino acid concentrations is linear and quite striking (Fig. 4.9). Whether or not S6 phosphorylation directly inhibits macroautophagy, just as is thought for S6 phosphorylation directly causing increased rates of protein synthesis, remains to be established. It seems more likely that the 70 kDa S6 kinase is also responsible for the phosphorylation of another protein which inhibits macroautophagy in its phosphorylated form. In addition to protein phosphorylation events regulating macroautophagy, intracellular phosphotidylinositols also appear to be important.20 Inhibitors of PI3 kinase inhibit macroautophagy under conditions in which hepatocyte cytoskeleton and ATP levels are not affected. In addition these inhibitors have no effect on cells when optimal amino acid suppression of macroautophagy has been achieved.20 However, PI3 kinase inhibitors have a variety of effects on cells, so a direct effect of phosphotidylinositols on macroautophagy remains to be proved.23 Several different research groups have implicated GTP-binding proteins in macroautophagy.19,30 In hepatocytes permeabilized with S. aureus α-toxin, formation and maturation of autophagosomes will occur if ATP and cytosol are supplied.19 However, these processes are blocked by the addition of small amounts of GTP-g−Σ. Many other

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Fig. 4.9. Inverse correlation between S6 phosphorylation and rates of proteolysis in rat liver. The varying rates of proteolysis were achieved by graded reductions in the levels of circulating amino acids. Proteolysis was measured as the release of radiolabeled valine previously incorporated into liver protein, and S6 phosphorylation was measured as incorporation of 32P into immunoprecipitated S6 protein.

forms of vesicular membrane traffic require small GTP-binding proteins in the ras-related GTPase (rab) family,68-70 and the inhibition of macroautophagy by GTP-g−S may be indicative of a role for rabs in macroautophagy. GTP may also regulate macroautophagy due to the involvement of a heterotrimeric G protein in this process.19,71 In the human colon cancer cell line, HT29, macroautophagic sequestration is dramatically inhibited by treatment with pertussis toxin which ADPribosylates and inactivates inhibitory G (Gi ) proteins. More impressively, overexpression of one isoform of Gi, Gi3, stimulates macroautophagy in proportion to its degree of overexpression.19 The mechanisms by which Gi3 stimulates macroautophagy remain to be discovered. In conclusion, regulators of macroautophagy already appear to be quite complex and to involve several different intracellular second messenger systems. The challenge for the future will be to discriminate between direct and indirect effects of inhibitors of second messenger pathways and to define the hierarchy of the signaling pathways.


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Substrate Selectivity As mentioned earlier, most studies of macroautophagy have emphasized its apparent nonselective capture of all cytoplasmic organelles and enzymes (Table 4.1). However, there are persistent and convincing reports of some selectivity in the uptake of proteins and organelles by macroautophagy under certain conditions. For example, peroxisomes can be induced to proliferate in rat liver due to treatment with clofibrate or other peroxisome proliferation agents. When the drug is removed from the hepatocytes, peroxisomes are selectively degraded by macroautophagy.72,73 In other experiments SER components were induced in rat liver by administration of phenobarbitol. When the drug was withdrawn, SER proteins were preferentially localized to autophagosomes and subsequently degraded in autolysosomes, presumably by macroautophagy.74 Insulin and amino acids reduce rates of protein degradation in rat liver and also correspondingly reduce rates of RNA degradation.3,10 The RNA that is degraded with suboptimal insulin and amino acid levels is primarily ribosomal RNA. However, glucagon treatment increases the rate of protein degradation without influencing the rate of RNA degradation.3,75 How ribosomes are excluded from macroautophagy under this condition remains a mystery. Selectivity in macroautophagic degradation has also been demonstrated in methylotrophic yeast.1,76 Such yeast grown on methanol contain abundant peroxisomes, organelles required for the metabolism of methanol. When such yeast are switched to ethanol as a carbon source, peroxisomes are first selectively sequestered into autophagosomes and then the autophagosomes fuse with the yeast vacuole. Other organelles, such as mitochondria, are excluded from this macroautophagy.76,77 The selective degradation of peroxisomes by macroautophagy also occurs in another methylotrophic yeast, Pichia methanolica.78 Macroautophagy was blocked at an early stage by mutations in acetyl coenzyme A synthase and in isocitrate lyase. The authors speculate that mutations in these enzymes block production of an ethanol metabolite that acts as a signal for the selective degradation of peroxisomes.78 This metabolite may be glyoxylate, but additional studies are required to prove this point. Additional insights into both selective and nonselective modes of macroautophagy come from studies of how the vacuolar protein, aminopeptidase I (API), is targeted from the cytosol, where it is synthesized, to the vacuole, where it resides.77 Mutations that fail to target API to the vacuole were isolated as cytoplasm-to-vacuole (cvt) mutants and compared with the apg macroautophagy mutants. Many, but not all, mutations were defective in both API import and in macroautophagy. These results indicate that the API import pathway partially overlaps with macroautophagic pathways. API is targeted to the vacuole after selective incorporation into small vesicles surrounded by a double membrane when yeast are grown in rich medium (Fig. 4.10).29 When yeast are deprived of nitrogen, API is delivered to the vacuole along with other cellular components by nonselective macroautophagy (Fig. 4.10).29 One interpretation of these results is that macroautophagy selective for API (and perhaps a few other vacuolar proteins targeted by the same mechanism) is responsible for delivery under optimal conditions while nonselective macroautophagy is responsible under deprivation conditions. Further studies along these lines may clarify how macroautophagy is able to operate in both selective and nonselective modes. Another protein conjugation system reminiscent of ubiquitination is required for the formation of autophagosomes.79 Agp7p has homology to the ubiquitin-activating enzyme (E1), and it is required to covalently link Agp5p to Apg12p.79-82


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Table 4.1. Degradation rates and lysosomal sequestration rates of 7 rat liver cytosolic enzymes Enzyme

Degradation rate (%/hr)

Sequestration rate (%/hr)

aldolase

3.9

3.6

lactate dehydrogenase

4.0

3.6

glucokinase

5.3

3.4

serine dehydratase

6.5

3.1

tryptophan oxygenase

19.0

3.7

tyrosine aminotransferase

20.0

3.5

ornithine decarboxylase

53.0

3.3

All proteins are sequestered into lysosomes at equivalent rates. Lysosomal sequestration can account for all of the degradation of some proteins (aldolase and lactate dehydrogenase) but can account for only 10% of the degradation of others (ornithine decarboxylase).

Future Directions of Research There are many outstanding problems regarding macroautophagy including the important issue of how autophagic vacuole membranes are formed. Identification of the membrane protein components has not lead to clarification of the origin of these membranes. Perhaps the permeabilized hepatocytes can be used to study the mechanisms of autophagosome formation including the origin of the autophagosome membrane. Identification of distinctive lipids in the autophagosome membrane may also be informative. The role of ubiquitination in the maturation of autophagic vacuoles requires additional experiments. Whether or not ubiquitination is required for the activity of components of the macrautophagic pathway, for optimal digestion of protein substrates within lysosomes, or a combination of both factors will help explain the link between ubiquitin and lysosomal proteolysis. Further studies using permeabilized hepatocytes may define roles for rabs and specific SNAREs in the fusion of late autophagosomes and lysosomes. For example, antibodies to a particular rab family member may block fusion of autophagosomes with lysosomes, while antibodies to another rab family member may prevent the initial formation of autophagosomes.The regulation of macroautophagy by amino acids has been widely studied, but the mechanisms of this regulation are not clear. Further studies aimed at the putative leucine receptor in the plasma membrane may lead to important molecular clues about regulation of macroautophagy. The second messengers that regulate macroautophagy are already numerous, but additional studies in this area that do not rely on inhibitors are important. The most convincing studies may use the yeast genetics together with biochemical studies to prove the direct effect of the mutations. Finally, the genetic analysis of the cvt pathway and the growing understanding of this vacuolar delivery mechanism promises to yield great insight about mechanisms of macroautophagy. Cvt and apg mutants that are in the same complementation groups will be interesting to define. Perhaps even more informative are the cvt and apg mutants that are defective only for cvt or apg.


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Fig. 4.10. Two pathways for the targeting of aminopeptidase I (API) from the cytosol to the yeast vacuole. During starvation API is targeted to the vacuole by way of the autophagosome. During growth the API is targeted to lysosomes by way of another organelle with a double membrane that selectively contains API and perhaps other vacuolar proteins that enter the vacuole by this mechanism.

References 1. Dunn WA. Autophagy and related mechanisms of lysosome-mediated protein degradation. Trends Cell Biol 1994; 4:139-143. 2. Seglen PO. Regulation of autophagic protein degradation in isolated liver cells. In: Glaumann H, Ballard FJ, eds. Lysosomes: Their Role in Protein Breakdown. New York: Academic Press, 1987:371-414. 3. Mortimore GE, Kadowaki M. Autophagy: Its mechanism and regulation. In: Ciechanover AJ, Schwartz AL, eds. Cellular Proteolytic Systems. New York: Wiley-Liss, 1994:65-87 4. Pfeifer U. Functional morphology of the lysosomal apparatus. In: Glaumann H, Ballard FJ, eds. Lysosomes: Their role in protein breakdown. New York: Academic Press, 1987:3-59. 5. Kominami E, Hashida S, Khairallah EA et al.. Sequestration of cytoplasmic enzymes in an autophagic vacuole-lysosome system induced by injection of leupeptin. J Biol Chem 1983; 258:6093-6100. 6. Kopitz J, Kisen G, Gordon PB et al. Nonselective autophagy of cytosolic enzymes in isolated rat hepatocytes. J Cell Biol 1990; 111:941-953. 7. Novikoff AB, Shin WY. Endoplasmic reticulum and autophagy in rat hepatocytes. Proc Nat Acad Sci USA 1978; 75:5039-5042.

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8. Dunn WA. Studies on the mechanism of autophagy: Formation of the autophagic vacuole. J Cell Biol 1990; 110:1923-1933. 9. Ueno T, Muno D, Kominami E. Membrane markers of endoplasmic reticulum preserved in autophagic vacuolar membranes from leupeptin-administered rat liver. J Biol Chem 1991; 266:18995-18999. 10. Mortimore GE, Pösö AR. Lysosomal pathways in hepatic protein degradation: Regulatory role of amino acids. Fed Proc 1984; 43:1289-1294. 11. Kovács AL, Grinde B, Seglen PO. Inhibition of autophagic vacuole formation and protein degradation by amino acids in isolated hepatocytes. Exp Cell Res 1981; 133:431-436. 12. Ballard FJ, Gunn JM. Nutritional and hormonal effects on intracellular protein catabolism. Nutr Rev 1982; 40:33-42. 13. Cockle SM, Dean RT. The regulation of proteolysis in normal fibroblasts as they approach confluence: Evidence for participation of the lysosomal system. Biochem J 1982; 28:795-800. 14. Knecht E, Hernandez-Yago J, Grisolia S. Regulation of lysosomal autophagy in transformed and nontransformed mouse fibroblasts under several growth conditions. Exp Cell Res 1984; 154:224-232. 15. Tanner A, Shen B-H, and Dice JF. Turnover of F1F0-ATP synthase subunit 9 and other proteolipids in normal and Batten disease fibroblasts. Biochim Biophys Acta 1997; 1361:251-262. 16. Holen I, Gordon PB, Seglen PO. Protein kinase-dependent effects of okadaic acid on hepatocyte autophagy and cytoskeletal integrity. Biochem J 1992; 284:633-636. 17. Falconer IR, Yeung DS. Cytoskeletal changes in hepatocytes induced by microcystis toxins and their relation to hyperphosphorylation of cell proteins. Chem Biol Interactions 1992; 81:181-196. 18. Blommaart EFC, Luiken JJFP, Blommaart PJE et al. Phosphorylation of ribosomal protein S6 is inhibitory for autophagy in isolated rat hepatocytes. J Biol Chem 1995; 270:2320-2326. 19. Ogier-Denis E, Couvineau A, Maoret JJ et al. A heterotrimeric Gi3 protein controls autophagic sequestration in the human colon cancer cell line HT-29. J Biol Chem 1995; 270:13-16. 20. Blommaart EFC, Krause U, Schellens JPM et al. The phosphotidylinositol 3-kinase inhibitors Wortmannin and LY294002 inhibit autophagy in isolated rat hepatocytes. A mechanism for 3-methyladenine action. Eur J Biochem 1997; 243:240-246. 21. Seglen PO, Gordon PB. 3-Methyladenine: Specific inhibitor of autophagic/lysosomal protein degradation in isolated hepatocytes. Proc Nat Acad Sci USA 1982; 79:1889-1892. 22. Hendil KB, Gordon PB, Seglen PO. Both endocytic and endogenous protein degradation in fibroblasts is stimulated by serum/amino acid deprivation and inhibited by 3-methyladenine. Biochem J 1990; 272:577-581. 23. de Camilli P, Emr S, McPherson PS et al. Phosphoinositides as regulators of membrane traffic. Science 1996; 271:1533-1539. 24. Gordon PB, Seglen PO. Autophagic sequestration of [14C]sucrose introduced into isolated rat hepatocytes by electropermeabilization. Exp Cell Res 1982; 142:1-14. 25. Tsukada M, Ohsumi Y. Isolation and characterization of autophagy-defective mutants of Saccharomyces cerevisiae. FEBS Lett 1993; 333:169-174. 26. Titorenko VI, Keizer J, Harder W et al. Isolation and characterization of mutants impaired in the selective degradation of peroxisomes in the yeast Hansenula polymorpha. J Bacteriol 1995; 177:357-363. 27. Matsuura A, Tsukada M, Wada Y et al. Apg1p, a novel protein kinase required for the autophagic process in Saccharomyces cerevisiae. Gene 1997; 192:245-250. 28. Funakoshi T, Matsuura A, Noda T et al. Analysis of APG13 gene involved in autophagy in yeast, Saccharomyces cerevisiae. Gene 1997; 192:207-213. 29. Baba M, Osumi M, Scott SV et al. Two distinct pathways for targeting proteins from the cytoplasm to the vacuole/lysosome. J Cell Biol 1997; 139:1687-1695. 30. Kadawaki M, Venerando R, Miotto G et al. De novo autophagic vacuole formation in hepatocytes permeabilized by Staphylococcus aureus α-toxin. Inhibition by GTP analogs. J Biol Chem 1994; 269:3703-3710. 31. Rabkin R, Tsao T, Shi JD et al. Amino acids regulate kidney cell protein breakdown. J Lab Clin Med 1991; 117:505-513. 32. Chua BH. Specificity of leucine effect on protein degradation in perfused rat heart. J Mol Cell Cardiol 1994; 26:743-751. 33. Rennie MJ, Hundal HS, Babij P et al. Characteristics of a glutamine carrier in skeletal muscle have important consequences for nitrogen loss in injury, infection, and chronic disease. Lancet 1986; 2:1008-1012.


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34. Dunlop DS, McHale DM, Lajtha A. The rate of protein degradation in developing brain. Methodological considerations. Biochem J 1982; 208:659-666. 35. Ahlberg J, Berkenstam A, Henell F et al. Degradation of short- and long-lived proteins in isolated rat liver lysosomes. Effects of pH, temperature, and proteolytic inhibitors. J Biol Chem 1985; 260:5847-5854. 36. Rock KL, Gramm C, Rothstein M et al. Inhibitors of the proteasome block the degradation of most cell proteins and the generation of peptides presented on MHC class I molecules. Cell 1994; 78:761-771. 37. Seglen PO, Gordon PB, Holen I. Nonselective autophagy. Semin Cell Biol 1990; 1:441-448. 38. Purhonen P, Persainen K, Reunanen H. Effects of brefeldin A on autophagy in cultured rat fibroblasts. Eur J Cell Biol 1997; 74:63-67. 39. Baba M, Ohsumi Y. Analysis of the membrane structure involved in autophagy in yeast by freezereplica method. Cell Struc Func 1995; 20:465-472. 40. Ueno T, Kominami E. Biochemical characterization of autolysosome membranes from leupeptinadministered rat liver. In: Ohsumi Y, ed. Dynamic Aspects of Lysosomal/Vacuolar System. Okazaki, Japan: Nat Inst Basic Biol, 1997:43. 41. Lenk SE, Dunn WA, Trausch JS et al. Ubiquitin-activating enzyme, E1, is associated with maturation of autophagic vacuoles. J Cell Biol 1992; 118:301-308. 42. Schwartz AL, Ciechanover A, Brandt RA et al. Immunoelectron microscopic localization of ubiquitin in hepatoma cells. EMBO J 1988; 7:2961-2966. 43. Laszlo L, Doherty FJ, Osborn NU et al. Ubiquitinated protein conjugates are specifically enriched in the lysosomal system of fibroblasts. FEBS Lett 1990; 261:365-368. 44. Gropper R, Brandt RA, Elias S et al. The ubiquitin-activating enzyme, E1, is required for stressinduced lysosomal degradation of cellular proteins. J Biol Chem 1991; 266:3602-3610. 45. Schworer CM, Schiffer HA, Mortimore GE. Quantitative relationship between autophagy and proteolysis during graded amino acid deprivation in perfused rat liver. J Biol Chem 1981; 256:7652-7658. 46. Seglen PO, Gordon PB. Amino acid control of autophagic sequestration and protein degradation in isolated rat hepatocytes. J Cell Biol 1984; 99:435-444. 47. Hoyvick H, Gordon PB, Berg TO et al. Inhibition of autophagic-lysosomal delivery and autophagic lactolysis by asparagine. J Cell Biol 1991; 113:1305-1312. 48. Mortimore GE, Pösö AR, Kadowaki M et al. Multiphasic control of hepatic protein degradation by regulatory amino acids. General features and hormonal modulation. J Biol Chem 1987; 262:16322-16327. 49. Miotto G, Venerando R, Siliprandi N. Inhibitory action of isovaleryl-L-carnitine on proteolysis in perfused rat liver. Biochem Biophys Res Commun 1989; 158:797-802. 50. Miotto G, Venerando R, Marin O et al. Inhibition of macroautophagy and proteolysis in the isolated rat hepatocyte by a nontransportable derivative of the multiple antigen peptide Leu8-Lys4Lys2 Lys-β-Ala. J Biol Chem 1994; 269:25348-25353. 51. Mortimore GE, Wert JJ, Miotto G et al. Leucine-specific binding of photoreactive Leu7-MAP to a high molecular weight protein on the plasma membrane of the isolated rat hepatocyte. Biochem Biophys Res Commun 1994; 203:200-208. 52. Blommaart EFC. Regulation of Hepatic Autophagy by Amino Acid Dependent Signal Transduction. PhD thesis, Biochemistry Department, Academic Medical Center, University of Amsterdam, The Netherlands, 1997. 53. Mortimore GE, Pösö AR, Lardeux BR. Mechanism and regulation of protein degradation in liver. Diabetes/Metab Rev 1989; 5:49-70. 54. Deter RL, Baudhuin P, de Duve C. Participation of lysosomes in cellular autophagy induced in rat liver by glucagon. J Cell Biol 1967; 35:c11-c15. 55. Woodside KH, Ward WF, Mortimore GE. Effects of glucagon on general protein degradation and synthesis in perfused rat liver. J Biol Chem 1974; 249:5458-5463. 56. Gilman AG. Guanine nucleotide-binding regulatory proteins and dual control of adenylate cyclase. J Clin Invest 1984; 73:1- 4. 57. Rosa F. Ultrastructural changes produced by glucagon, cyclic 3':5'-AMP, and epinephrine in perfused livers. J Ultrastruct Res 1971; 34:205-213. 58. Aplin A, Jasionowski T, Tuttle DL et al. Cytoskeletal elements are required for the formation and maturation of autophagic vacuoles. J Cell Physiol 1992; 152:458-466.

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59. Fengsrud M, Roos N, Berg T et al. Ultrastructural and immunocytochemical characterization of autophagic vacuoles in isolated hepatocytes: effects of vinblastine and asparagine on vacuole distribution. Exp Cell Res 1995; 221:504-519. 60. Collot M, Louvard D, Singer SJ. Lysosomes are associated with microtubules and not with intermediate filaments in cultured fibroblasts. Proc Nat Acad Sci USA 1984; 81:788-792. 61. Häussinger D, Hallbrucker C, Vom Dahl S et al. Cell swelling inhibits proteolysis in perfused rat liver. Biochem J 1990; 272:239-242. 62. Häussinger D, Roth E, Lang F et al. Cellular hydration state: an important determinant of protein catabolism in health and disease. Lancet 1993; 341:1330-1332. 63. Hallbrucker C, Vom Dahl S, Lang F et al. Control of hepatic proteolysis by amino acids. The role of cell volume. Eur J Biochem 1991; 197:717-724. 64. Garrison JC. The effects of glucagon, catecholamines, and the calcium ionophore A23187 on the phosphorylation of rat hepatocyte cytosolic proteins. J Biol Chem 1978; 253:7091-7100. 65. Holen I, Strømhaug PE, Gordon PB et al. Inhibition of autophagy and multiple steps in asialoglycoprotein endocytosis by inhibitors of tyrosine protein kinases (tyrphostins). J Biol Chem 1995; 270:12823-12831. 66. Gordon PB, Holen I, Fosse M et al. Dependence of hepatic autophagy on intracellularly sequestered calcium. J Biol Chem 1993; 268:26107-26112. 67. Blommaart EFC, Luiken JJFP, Meijer AJ. Autophagic proteolysis: Control and specificity. Histochem J 1997; 29:365-385. 68. Goud B, McCaffrey M. Small GTP-binding proteins and their role in transport. Curr Opin Cell Biol 1991; 3:626-633. 69. Balch WE. From G minor to G major. Curr Biol 1992; 2:157-160. 70. Melancon P. Vesicle traffic: “G whizz”. Curr Biol 1993; 3:230-233. 71. Huang C, Hepler JR, Chen LT et al. Organization of G proteins and adenyl cyclase at the plasma membrane. Mol Biol Cell 1997; 8:2365-2378. 72. Luiken JJFP, van den Berg M, Heikoop JC et al. Autophagic degradation of peroxisomes in isolated rat hepatocytes. FEBS Lett 1992; 304:93-97. 73. Yokota S. Formation of autophagosomes during degradation of excess peroxisomes induced by administration of dioctyl phthalate. Eur J Cell Biol 1993; 61:67-80. 74. Masaki R, Yamamoto A, Tashiro Y. Cytochrome P-450 and NADPH-cytochrome P-450 reductase are degraded in the autolysosomes in rat liver. J Cell Biol 1987; 104:1207-1215. 75. Lardeux BR, Mortimore GE. Amino acid and hormonal control of macromolecular turnover in perfused rat liver. Evidence for selective autophagy. J Biol Chem 1987; 262:14514-14519. 76. Tuttle DL, Dunn WA. Divergent modes of autophagy in the methylotrophic yeast Pichia pastoris. J Cell Sci 1995; 108:25-35. 77. Klionsky DJ. Protein transport from the cytoplasm into the vacuole. J Membr Biol 1997; 157:105-115. 78. Kulachkovsky AR, Moroz OM, Sibirny AA. Impairment of peroxisome degradation in Pichia methanolica mutants defective in acetyl CoA synthetase or isocitrate lyase. Yeast 1997; 13:1043-1052. 79. Mizushima N, Noda T, Yoshimori T et al. A protein conjugation system essential for autophagy. Nature 1998; 395:395-398. 80. Kim J, Dalton VM, Eggerton KP et al. Apg7p/Cvt2p is required for the cytoplasm-to-vacuole targeting, macroautophagy, and peroxisome degradation pathways. Mol Biol Cell 1999; 10:13370-1351. 81. Tanida I, Mizushima N, Kiyooka M et al. Apg7p/Cvt2p: A novel protein activating enzyme essential for autophagy. Mol Biol Cell 1999; 10:1367-1379. 82. Yuan W, Stromhaug PE, and Dunn W. Glucose-induced autophagy of peroxisomes in Pichia pastoris requires a unique #1-like protein. Mol Biol Cell 1999; 10:1353-1366. 83. Branch AD. A good antisense molecule is hard to find. Trends Biochem Sci 1998; 23:45-50.

CHAPTER 5

Degradation of Intracellular Proteins by Microautophagy Overview

M

icroautophagy was first proposed by de Duve and Wattiaux1 more than 30 years ago. This term referred to the then hypothetical notion that smaller bits of cytoplasm, perhaps even below the limits of detection by electron microscopy, could be taken up by lysosomes in addition to the more morphologically obvious macroautophagy. Some of the confusion in the literature regarding microautophagy results from the same name being applied to what may be very different lysosomal processes.2-5 Microautophagy refers to the internalization of bits of cytoplasm by vesicle formation at the lysosome membrane. The size of the vesicle and its mechanisms of formation appear to be variable (Fig. 5.1).2,3 Small organelles such as ribosomes and glycogen particles can be seen in these vesicles, but larger organelles such as mitochondria are usually excluded. Ahlberg and Glaumann2,3 described the in vitro uptake of particles and proteins by isolated lysosomes as reflections of microautophagy that result in multivesicular bodies (Fig. 5.2). However, large organelles such as peroxisomes can also be taken up by the yeast vacuole in a process also called microautophagy6 that involves wrapping of the peroxisome in a flaplike protrusion from the vacuole membrane (Figs. 5.1C and 5.3). Finally, a different shape of some lysosomes in rat liver has been proposed to reflect microautophagy,7,8 but these lysosomes appeared to contain cup-like contents (Figs. 5.1B and 5.4) rather than the multivesicular body appearance. Microautophagy has also been defined biochemically as the lysosomal component of protein degradation that occurs under conditions in which macroautophagy is maximally suppressed.4,9,10 For example, in the perfused rat liver macroautophagy can be inhibited by including amino acids and insulin in the perfusion medium.11,12 Under these conditions, lysosomes still contain some substrate proteins that have presumably entered lysosomes by mechanisms other than macroautophagy. However, these proteins may have entered the lysosome by the selective, chaperone-mediated pathway (see Chapter 6) in addition to microautophagy.

Methods Used to Study Microautophagy The tissue in which microautophagy has been most widely studied is the liver, but microautophagy undoubtedly occurs to some extent in all lysosomes and in the yeast homologous structure, the vacuole. As mentioned previously, the electron microscopic


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Fig. 5.1. Different forms of microautophagy. (A) Small vesicles form at the lysosome (L) membrane, and these small vesicles are internalized by the lysosome to give the appearance of a multivesicular body (MVB). (B) Larger vesicles may also form at the lysosomal membrane. (C) Lysosomes may also extend a portion of their membrane to surround a bit of cytoplasm and generate a “padlock”structure.

appearance of lysosomes that are active in microautophagy may be variable, perhaps reflecting the size of the internalized lysosomal membrane area. The biochemical definition of microautophagy, the lysosomal component to the degradation of long-lived proteins when macroautophagy has been maximally suppressed,4,9,10 has been useful for the study of possible regulation of this process. However, as mentioned above, it is possible that protein substrates which enter lysosomes under these conditions are doing so by more than one pathway. The ability to reproduce microautophagy using isolated lysosomes has also contributed to our understanding of this process.2,3 Rats were given injections of iron to increase the density of residual bodies, and then lysosomes were separated from residual bodies over Percoll density gradients. The lysosomes, but not the residual bodies, were able to take up and degrade proteins and were also able to internalize electron dense Percoll beads and ferritin particles (Figs. 5.2 and 5.5).3 Microsomes (sealed vesicles derived from the endoplasmic reticulum) incubated in vitro showed none of this activity. These in vitro studies

Degradation of Intracellular Proteins by Microautophagy

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Fig. 5.2. Rat liver lysosomes incubated in vitro in the presence of electron dense substances such as Percoll beads or ferritin. (A) Percoll beads or ferritin can be found inside lysosomes in what appear to be small vesicles after a 20 min incubation. (B)The membrane of the small vesicles eventually breaks down, and the Percoll beads in the lysosome are no longer contained within vesicles after an additional 30 min incubation.

showed that degradation of proteins was ATP-dependent, but the formation of microautophagic vesicles was not.2 Recent studies have shown that microautophagy can also take place in yeast. When methylotrophic yeast such as Pichia pastoris are grown in methanol, they induce peroxisomes and cytosolic enzymes required to utilize this carbon source. When these yeast are switched to a glucose-containing medium, the peroxisomes are degraded within the vacuole by a process resembling microautophagy. That is, peroxisomes are surrounded by flap-like extensions of the vacuolar membrane and enter the vacuole in a vesicle derived from the lysosomal membrane (Fig 5.3).6,13 Isolation of mutants defective in microautophagy should provide clues about microautophagic mechanisms. A second methylotrophic yeast, Hansenula polymorpha, shows the same rapid degradation of peroxisomes when placed in glucose-containing medium. However, the peroxisomes are delivered to vacuoles after being surrounded by a double membrane in a process similar to macroautophagy.14 Recall that when this yeast type is changed from methanol to ethanol, peroxisomes are also degraded but by a process resembling macroautophagy (see Chapter 4).

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Fig. 5.3. A flap-like extension from the lysosome can surround an organelle. (A) A lysosome (L) deforms to surround a peroxisome (P). (B) The protrusion extends further and eventually fuses with another region of the lysosome membrane. (C) This process leads to peroxisomes contained within the vacuole surrounded, at least transiently, by two membranes. The peroxisomal membrane presumably breaks down, and the entire content of the peroxisome is degraded.

A recent study of microautophagy in the yeast, Pichia pastoris, utilized double fluorescent markers to label peroxisomes and the vacuolar membrane. Peroxisomes were tagged with green fluorescent protein containing a carboxyl terminal SKL tripeptide sequence, a type 1 peroxisomal targeting peptide.15,16 Vacuolar membranes were fluorescently labeled with the dye, FM4-64. Morphological analysis of peroxisomes and the vacuolar membrane after switching carbon source from methanol to glucose revealed several intermediate steps in the microautophagic process (see below).

Mechanisms of Microautophagy Little is known with certainty regarding the mechanisms or regulation of microautophagy. The vesiculation of the lysosome surface in vitro does not appear to


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Fig. 5.4. Morphological appearance of type A and type R lysosomes in mouse liver. The type A lysosomes have a “cup-like” appearance of their contents and could reflect microautophagy of a single, large vesicle. Type R lysosomes have a uniform, electron dense appearance.

require ATP, but the subsequent degradation of vesicles and their contents does require ATP2 at least for the maintenance of the acidic conditions within the lysosome, but possibly for other reasons as well. The forces causing the lysosomal membrane to vesiculate are completely unknown. The intralysosomal vesicles do not appear to be coated with protein, and nothing is known about their lipid composition. An intact cytoskeleton is not required for microautophagy,2,3 but microautophagy of peroxisomes by Pichia pastoris does require ongoing protein synthesis.6,13 A short-lived protein may be a necessary component of this process. An ATP-independent vesiculation at the plasma membrane has been shown to be induced by sphingomyelinase treatment.17 An acidic form of this enzyme is normally located in lysosomes and endosomes, and a neutral form is in the cytosol.18 Perhaps sphingomyelinase action or the subsequent release of cholesterol from the treated membrane may be responsible for inward vesicle formation from the lysosomal membrane. There may be some selectivity in the microautophagic uptake of particular proteins. For example, Ahlberg and Glaumann2 showed that lysosomes in vitro degraded methemoglobin 5-times faster than lysozyme and 10-times faster than ovalbumin (Fig. 5.5). However, some or all of this difference could be caused by differences in the protein's susceptibility to proteolytic attack once inside the lysosome. Alternatively, proteins may somehow be able to bind to the lysosomal membrane region about to be internalized, and methemoglobin may bind more avidly than lysozyme or ovalbumin. The recent detailed study of microautophagy in Pichia pastoris mentioned earlier using double fluorescent labeling of peroxisomes and the vacuole membrane identified several

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Fig. 5.5. Degradation of three proteins added to isolated intact rat liver lysosomes. The degradation rates were measured as the conversion of acid-precipitable radioactivity to acid-soluble radioactivity. The lysosomes were also incubated with electron dense ferritin, and ferritin could be seen by electron microscopy to be contained within intralysosomal vesicles (see Figure 5.2). The more rapid degradation of [14C]methemoglobin is consistent with some selectivity in microautophagic internalization and/or differences in substrate susceptibility to proteolytic attack once inside lysosomes.

intermediates in the microautophagy of peroxisomes19 (Fig. 5.6). When the yeast are growing in methanol, the vacuole and clusters of peroxisomes are distinct structures within cells, and there are few, if any, invaginations or protrusions from the vacuole membrane (stage 0). After switching to glucose as a carbon source, the vacuolar membrane can be seen to invaginate with clusters of peroxisomes at the point of invagination (early stage 1). These invaginations became deeper and the membrane structure became more complex (late stage 1). The intact clusters of peroxisomes can then be seen within a sealed vesicle within the vacuole (stage 2). The green fluorescent protein-SKL localized to peroxisomes then can be found within the intravacuolar vesicle and then throughout the vacuole (stage 3). Eventually, all of the green fluorescent protein-SKL was degraded, and the vacuoles were no longer fluorescent (stage 4).19 As mentioned earlier, cycloheximide inhibits microautophagy. In the presence of cycloheximide, peroxisome clusters near shallow invaginations in the vacuolar membrane could be seen, but deep invaginations were absent. These results indicate that ongoing protein synthesis is required for the progression from early stage 1 to late stage 1 (Fig. 5.6).19 Both microautophagy and macroautophagy are reduced in yeast that are deficient in the vacuolar proteases A and B. When following macroautophagic degradation of peroxisomes after switching from methanol to ethanol, there was no defect in the delivery of peroxisomes to the vacuole, but there was reduced degradation of the peroxisomes once inside the vacuole. In contrast, a deficiency of proteases A and B blocked microautophagy between stage 0 and stage 1. In other words, proteases A and B appear to be required for the invagination of the vacuolar membrane at the site of the peroxisome clusters (Fig. 5.6).17


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Fig. 5.6. Stages of microautophagy in Pichia pastoris. Stage 0: Clusters of peroxisomes (P) are distinct from the vacuole (V). Stage 1, early: Clusters of peroxisomes are localized to sites of indentation of the vacuole membrane. Stage 1, late: Clusters of peroxisomes are localized to deep indentations of the vacuolar membrane. Stage 2: Peroxisomes are contained in a vesicle within the vacuole. Stage 3: The peroxisomes begin to be degraded as does the vesicle membrane within the vacuole. Peroxisomal matrix proteins can be seen throughout the vacuole. Stage 4: All peroxisomal and vesicle proteins are degraded. The stages at which cycloheximide, phenylmethyl-sulfonylfluoride (PMSF), an inhibitor of proteinases A and B, and various pag mutants are blocked are indicated.

Several mutants that were defective in the degradation of green fluorescent protein-SKL were isolated (deficient in peroxisome degradation by microautophagy; pag) and divided into 6 different complementation groups. Morphological analysis of these mutants support the 4 stage model of microautophagic degradation of peroxisomes (Fig. 5.6). Pag1 and

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pag2 are deficient between stages 0 and 1. No invaginations of the vacuolar membrane were evident in these mutants. Pag3 mutants showed deep invaginations of the vacuolar membrane that contained peroxisome clusters, but the invaginations did not close and the peroxisomes were not degraded. Pag4, pag5, and pag6 mutants resulted in peroxisomes residing within vesicles in the vacuole, but the peroxisomes were not degraded. Therefore, the block in these mutants appeared to be between stages 2 and 3. Importantly, none of the pag mutants affected macroautophagic degradation of peroxisomes in response to a shift from methanol to ethanol.19 One mutation in Pichia pastoris that is defective in microautophagy of peroxisomes has been analyzed, and the mutation surprisingly lies in the gene encoding the α subunit of phosphofructokinase.20 The requirement for phosphofructokinase in allowing microautophagy to occur did not depend on its known enzymatic activity, suggesting that carbohydrate metabolism was not involved. It is possible that phosphofructokinase is a protein with multiple functions in these yeast,20 but its exact role in microautophagy remains unknown. Interestingly, the same protein conjugation system required for macroautophagy is also required for microautophagy.21

Regulation of Microautophagy The biochemical definition of microautophagy as the protein substrates that enter lysosomes when macroautophagy is maximally suppressed indicates, by definition, that microautophagy is not affected by amino acids or insulin. However, Mortimore and colleagues identified two kinds of morphologically distinct lysosomes, type A and type R (Fig. 5.4) that may indicate two different forms the microautophagic process. The volume of type A lysosomes increased during 48 hrs of starvation in the mouse, but the volume of type R lysosomes were not affected by starvation (Fig. 5.7). Pharmacological doses of glucagon also increase the number of type A lysosomes, but the physiological significance of this action is not known.4

Future Directions of Research Analysis of the pag mutants identified in Pichia pastoris will be very important in better defining the mechanisms and regulation of microautophagy. These mutations will probably include defective structural components of microautophagy and perhaps regulatory machinery as well. Secondary effects of the mutations can be best demonstrated using in vitro reconstitution of the microautophagic process. If such systems are not available, the proteins encoded by the pag mutants will have to be identified. The genes can be placed under the control of a repressor, or temperature-sensitive mutations can be isolated, and the timing of the mutation on microautophagy examined. A primary effect should be evident soon after repression of synthesis if the protein is short-lived and soon after shift to the nonpermissive temperature in the case of the temperature-sensitive mutations. Whether or not lysosomal or cytosolic sphingomyelinase is required for microautophagy might be addressed using fibroblasts from patients with Niemann-Pick type A disease that lack lysosomal sphingomyelinase. A defect in microautophagy could most easily be detected using isolated lysosomes and following their ability to take up electron dense tracers such as ferritin. A role for cytosolic sphingomyelinase might also be assessed with this system. Some microautophagy occurs in isolated lysosomes without added sphingomyelinase, but some of the enzyme might be present in the lysosomal fraction. A


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Fig. 5.7. Relative volumes of type R and type A secondary lysosomes in mouse liver under conditions of feeding and fasting. Volumes were plotted against measured rates of proteolysis. It is not clear whether or not either of these types of lysosomes corresponds to microautophagy.

decrease in microautophagy in the presence of neutralizing antibodies to sphingomyelinase and an increase in microautophagy due to added sphingomyelinase might prove a role for this enzyme in the microautophagic process. It is also possible that some other lipidmodifying enzymes alter lysosomal lipids in a way that induces microautophagy. The possibility of selectivity in microautophagic protein uptake into lysosomes requires further research. If proteins such as methemoglobin are preferential substrates for microautophagy, it may show some degree of binding to lysosomal membranes. The nature of this binding could be explored with a variety of affinity assays, and the binding macromolecules could be characterized as protein, lipid, and/or carbohydrate. References 1. de Duve C, Wattiaux R. Functions of lysosomes. Ann Rev Physiol 1966; 28:435-492. 2. Ahlberg J, Glaumann H. Uptake-microautophagy and degradation of exogenous proteins by isolated rat liver lysosomes. Effects of pH, ATP, and inhibitors of proteolysis. Exp Mol Path 1985; 42:78-88. 3. Ahlberg J, Marzella L, Glaumann H. Uptake and degradation of proteins by isolated rat liver lysosomes. Suggestion of a microautophagic pathway of proteolysis. Lab Invest 1982; 47:523-532. 4. Mortimore GE, Kadowaki M Authophagy: Its mechanism and regulation. In: Ciechanover AJ, Schwartz AL, eds. Cellular Proteolytic Systems. New York: Wiley-Liss, 1994: 65-87. 5. Pfeifer U. Functional morphology of the lysosomal apparatus. In: Glaumann H, Ballard FJ, eds. Lysosomes: Their role in protein breakdown. New York: Academic Press, 1987:3-59.

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6. Dunn WA. Autophagy and related mechanisms of lysosome-mediated protein degradation. Trends Cell Biol 1994; 4:139-143. 7. Mortimore GE, Pösö AR, Lardeux BR. Mechanism and regulation of protein degradation in liver. Diabetes/Metab Rev 1989; 5:49-70. 8. Mortimore GE, Hutson NJ, Surmacz CA. Quantitative correlation between macro- and microautophagy in mouse hepatocytes during starvation and refeeding. Proc Nat Acad Sci USA 1983; 80:2179-2183. 9. Mortimore GE, Pösö AR. Lysosomal pathways in hepatic protein degradation: Regulatory roles for amino acids. Fed Proc 1984; 43:1289-1294. 10. Mortimore GE, Pösö AR. Intracellular protein catabolism and its control during nutrient deprivation and supply. Ann Rev Nutr 1987; 7:539-564. 11. Lardeux BR, Mortimore GE. Amino acid and hormonal control of macromolecular turnover in perfused rat liver. Evidence for selective autophagy. J Biol Chem 1987; 262:14514-14519. 12. Mortimore GE, Mondon CE. Inhibition by insulin of valine turnover in liver. J Biol Chem 1970; 245:2375-2383. 13. Tuttle DL, Dunn WA. Divergent modes of autophagy in the methylotrophic yeast Pichia pastoris. J Cell Sci 1995; 108:25-35. 14. Veenhuis M, Douma A, Harder W et al. Degradation and turnover of peroxisomes in the yeast Hansenula polymorpha induced by selective inactivation of peroxisomal enzymes. Arch Microbiol 1983; 134:193-203. 15. Subramani S. Protein import into peroxisomes and biogenesis of the organelle. Ann Rev Cell Biol 1993; 9:445-498. 16. Subramani S. Components involved in peroxisome import, biogenesis, proliferation, turnover, and movement. Physiol Rev 1998; 78:1-18. 17. Zha X, Pierini LM, Leopold PL et al. Sphingomyelinase treatment induces ATP-independent endocytosis. J Cell Biol 1998; 140:39-47. 18. Hanun YA. The sphingomyelin cycle and the second messenger function of ceramide. J Biol Chem 1994; 269:3125-3128. 19. Sakai Y, Koller A, Rangell LK et al. Peroxisome degradation by microautophagy in Pichia pastoris. Identification of specific steps and morphological intermediates. J Cell Biol 1998; 141:625-636. 20. Yuan W, Tuttle DL, Ralph GS et al. Glucose-induced microautophagy in Pichia pastoris requires the α-subunit of phosphofructokinase. J Cell Sci 1997; 110:1935-1945. 21. Yuan W, Stromhaug PE, Dunn WA. Glucose-induced autophagy of peroxisomes in Pichia pastoris requires a unique E1-like protein. Mol Biol Cell 1999; 10:1353-1366.

CHAPTER 6

Selective Pathway for Degradation of Cystolic Proteins by Lysosomes Overview

L

ysosomes are able to take up certain cytosolic proteins in a molecule-by-molecule fashion. This selective pathway of lysosomal proteolysis resembles in many respects the movement of proteins into the lumen of the endoplasmic reticulum (ER) and the import of proteins into mitochondria, chloroplasts, and peroxisomes.1-4 Redundant but biochemically related targeting peptides are components of all known substrate proteins for these targeting pathways. The substrate proteins bind to receptors in the organelle membranes, and then, with or without unfolding, are transported through a gated aqueous pore. Molecular chaperones outside and inside the organelle are required for separate steps in the import pathways.1-4

Background

By the late 1960s it was clear that proteins had widely varying half-lives,5 some examples of which are listed in Table 6.1. Such variability in degradation rates could not be easily explained based on electron microscopic images of entire mitochondria, for example, being engulfed by autophagic vacuoles and then digested by lysosomal cathepsins. Several investigators made innovative proposals about how proteins might selectively enter lysosomes by macroautophagy or microautophagy6 or how certain proteins might escape the digestion within lysosomes and reenter the cytosolic compartment.7 Certain aspects of these proposals still have merit.8,9 We began studying the enhanced protein degradation seen in tissues of diabetic and starved rats.10,11 In the liver of acutely diabetic rats the evidence was quite clear that the rate of protein degradation was increased.12 Early in starvation and after acute insulin withdrawal from rats made diabetic by injections of streptozotocin, degradation rates of most proteins were increased. This period also coincided with an increased number and size of macroautophagic vacuoles.13,14 At later times of starvation or insulin withdrawal, particular classes of proteins were degraded at enhanced rates but other classes were not.15 For example, small, basic, nonglycosylated proteins were degraded more rapidly, but large, acidic, glycoproteins were not, and this caused the small, basic, nonglycosylated proteins to be preferentially lost from the liver cytosol.16 Withdrawal of insulin from hepatocytes in cell culture also caused increased proteolysis, and this increase could be blocked by inhibitors of lysosomal proteolysis.17,18 Although other explanations regarding our results with diabetic rats were possible, the simplest explanation was that lysosomes could operate Lysosomal Pathways of Protein Degradation, by J. Fred Dice, Ph.D. ©2000 EUREKAH.COM


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Table 6.1. Half-lives of different proteins in rat liver Protein

Half-Life (hrs)

ornithine decarboxylase

0.3

RNA polymerase I

1

cyclins

2-5

cAMP-dependent protein kinase

2-8

cGMP-dependent protein kinase

2-8

β-spectrin

8

glucokinase

11

catalase

30

histidase

60

lactate dehydrogenase

171

superoxide dismutase

186

histones

2800

Half-lives were determined by pulse-chase protocols followed by purification of the proteins. Original literature citations can be found in reference 4.

in a selective fashion at later times of starvation and insulin withdrawal. We now know that this is true and have been fortunate to have made progress in understanding this selective lysosomal pathway of proteolysis.

Methods Used for the Study of Selective Lysosomal Proteolysis We realized that studying protein degradation in whole animals limited our ability to analyze protein degradative pathways in molecular detail. We spent several months isolating hepatocytes from rats with the hope of studying protein degradation in these nondividing cells after they form a monolayer. Such hepatocyte monolayers were not responsive to insulin, however, probably due to damage of their insulin receptors during the hepatocyte isolation.19 We next turned to a readily available human lung fibroblast cell called IMR-90.20 These cells are primary, nontransformed cells that grow as a monolayer until they become confluent, after which time growth is markedly reduced. Confluent cultures could be maintained in tissue culture medium with or without added serum growth factors. In the absence of serum rates of proteolysis transiently double when compared to rates of proteolysis in cells maintained in the presence of 10% fetal or newborn calf serum (Fig. 6.1). The serum factors that contribute to reducing protein degradation include insulin, insulin-like growth factor, fibroblast growth factor, and a cortisol analog, dexamethasone.21 One reason we wanted a cell culture model to study protein degradation was that we wanted to be able to introduce a single radiolabeled protein into the cytosol of cells to

Selective Pathway for Degradation of Cystolic Proteins by Lysosomes

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Fig. 6.1. Degradation of long-lived proteins in fibroblasts maintained with (+S) and without (-S) 10% serum. Confluent cultures of IMR-90 fibroblasts were labeled for 2 days in the presence of serum with [3H]leucine and chased in medium containing excess unlabeled leucine. Loss of acid-precipitable radioactivity from the monolayers is shown. The numbers refer to half-lives in hrs.

follow its rate of degradation. The study of degradation of all cell proteins or entire molecular classes of proteins seemed inherently confusing because growing evidence was emerging that many pathways of proteolysis coexisted within cells.22,23 An emerging technique in the late 1970s pioneered by the laboratories of Okada,24 Loyter,25 and Rechsteiner26 was to load a radiolabeled protein into red cell ghosts, and to use these loaded ghosts to deliver the labeled protein to the cytosol of cultured cells. To accomplish this delivery, the red cell membrane must be induced to fuse with the cultured cell's plasma membrane. This fusion could be accomplished using inactivated viruses or their isolated fusion proteins or chemical fusogens such as polyethylene glycol (Fig. 6.2).24-26 Many cell types in culture can be successfully microinjected with only slight modifications of the experimental protocols.27 Under the right conditions cells fully recover from the microinjection procedures within 2 hours. Our initial studies with this technique used a mixture of cytosolic proteins derived from IMR-90 fibroblasts that we radiolabeled with 125I and then microinjected back into the cytosol of IMR-90 fibroblasts.28 This complex mixture of proteins was degraded with the same characteristics as the endogenous IMR-90 cytosolic proteins.28 These results demonstrated the validity of using this technique to study protein degradation. We and others also showed that the half-life of microinjected proteins was independent of the amount of protein injected indicating that the amount of protein injected was not saturating

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Fig. 6.2. Schematic representation of the steps in red cell-mediated microinjection 1. Red blood cells (RBC) are lysed in a hypotonic medium to remove most of the hemoglobin (Hb). 2. A radiolabeled protein (*) is added in hypotonic medium, and some of the radiolabeled protein enters the red cell ghost. 3. The mixture is returned to physiological saline, and the red cell ghost membrane reseals. 4. The uncaptured radiolabeled protein is removed by repeated centrifugation and collection of the red cell ghosts. 5. The “loaded” red cells are allowed to contact a monolayer of human fibroblasts, and membrane fusion is induced by addition of a fusogen. The radiolabeled protein diffuses into the cytosol of the recipient fibroblast.

any proteolytic pathways.28 Some of these individual microinjected proteins (ribonuclease A, aspartate aminotransferase) were degraded more rapidly in response to serum withdrawal while others (insulin A chain, lysozyme) were degraded at the same rate in the presence and absence of serum (Fig. 6.3).28 The enhanced degradation of several of the proteins in response to serum withdrawal could be partially blocked by inhibitors of lysosomal proteolysis such as NH4Cl.29 These results suggested that the enhanced protein degradation in response to serum withdrawal was lysosomal and was selective. To prove that the degradation of microinjected ribonuclease A (RNase A) was due to lysosomal proteolysis, we tagged the RNase A with a radiolabeled, nonmetabolizable sugar, [3H]raffinose.29 The [3H]raffinose was chemically linked to lysines in the RNase A and


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Fig. 6.3. Degradation rates of some microinjected proteins are increased following serum withdrawal. The rate of loss of microinjected radioactive proteins was followed in confluent cultures of IMR-90 human fibroblasts. Solid circles and lines represent degradation in the presence of serum, while open circles and dashed lines represent degradation in the absence of serum. RNase A = ribonuclease A; AAT = aspartate aminotransferase (cytosolic form); INS-A = insulin A chain; LYSOZ = lysozyme. The numbers refer to the half-lives in hrs.

the resultant compound was microinjected into fibroblasts. The degradation products, mainly [3H]raffinose-lysine, were found to accumulate entirely within lysosomes.29 In contrast, [3H]raffinose-lysine derived from microinjected [3H]raffinose-bovine serum albumin accumulated primarily in the cytosol. Interestingly, even though all of the degradation of RNase A was lysosomal, addition of NH4Cl inhibited its degradation by only 50%. We concluded that lysosomotropic agents underestimate the role of lysosomes in proteolysis probably because some cathepsins retain activity at pH 7.0.29

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The reproduction of this lysosomal pathway of proteolysis using highly purified lysosomes from human fibroblasts and from rat liver has greatly extended our understanding of the biochemical requirements for this pathway of proteolysis. For example, the role of the molecular chaperone, the constitutively expressed heat shock protein of 73 kilodaltons (hsc73), in stimulating lysosomal uptake of substrate proteins would have been difficult to address without the availability of in vitro systems because of the multiple roles of hsc73 in living cells.

RNase A as a Substrate for Selective Lysosomal Proteolysis We analyzed further the serum regulated degradation of RNaseA after radiolabeling the protein with 125I , 131I, or 3H. The method of labeling did not influence the results we obtained.28,30 RNase A (Fig. 6.4) can be cleaved by the protease subtilisin into the amino terminal 20 amino acids (RNase S-peptide) and the 21-124 remaining amino acids (RNase S-protein). After microinjection into human fibroblasts, RNase A was degraded more rapidly in response to serum withdrawal, but RNase S-protein was degraded at the same rate in the presence and absence of serum (Fig. 6.5).30 We obtained this result even when [125I]RNase A and [131I]RNase S-protein were co-loaded into red cell ghosts and comicroinjected into the very same fibroblasts.30 In addition, RNase S-peptide microinjected by itself was degraded in a serum-regulated manner (Fig. 6.5).24 These results indicated that the information required for RNase A to be degraded more rapidly in response to serum withdrawal was contained in its amino terminal 20 amino acids. By analyzing the ability of smaller synthetic peptides to inhibit the increased degradation of microinjected RNase A in response to serum withdrawal, we were able to identify residues 7-11 of RNase S-peptide, KFERQ, as the critical region for targeting the protein to lysosomes for enhanced degradation in response to serum withdrawal (Fig. 6.6).31 We showed that RNase S-peptide could be chemically linked to three unrelated proteins that did not exhibit serum-regulated degradation, and in each case the conjugate was now degraded in a serum-responsive fashion (Fig. 6.7).32 These studies showed the RNase S-peptide contained information that was both necessary and sufficient for targeting heterologous proteins for enhanced degradation in response to serum withdrawal.

The KFERQ Motif We noted that the exact peptide sequence, KFERQ, is found only in members of the RNase A family. However, experimental results indicated that biochemically related peptides are present in all substrates of this pathway of lysosomal proteolysis. We raised antibodies to KFERQ linked to a carrier protein, and these antibodies were able to immunoprecipitate 30% of cytosolic proteins.33 By radiolabeling cellular proteins and following their degradation in the presence and absence of serum, we could follow the degradation of immunoprecipitable and nonimmunoprecipitable proteins. The enhanced degradation in response to serum withdrawal applied only to the immunoprecipitable proteins.33 To identify such biochemically related peptide regions, we searched the primary sequences of 4 unrelated proteins that were known substrates for this selective lysosomal pathway of proteolysis and looked for sequences that were not present in four proteins that were known not to be substrates.33,34 Table 6.2 shows the putative peptides related to KFERQ. The consensus motif identified consisted of a basic amino acid, an acidic amino acid, a very hydrophobic amino acid, and a second basic or very hydrophobic amino acid in any order on either side of a required glutamine. Many additional substrates and nonsubstrates for the selective lysosomal proteolytic pathway have since been identified35-38


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Fig. 6.4. The amino acid sequence of RNase A. Single letter abbreviations for amino acids are used. A = alanine; C = cysteine; D = aspartate; E = glutamate; F = phenylalanine; G = glycine; H = histidine; I = isoleucine; K = lysine; L = leucine; M = methionine; N = asparagine; P = proline; Q = glutamine; R = arginine; S = serine; T = threonine; V = valine; Y = tyrosine. The arrow shows the cleavage site of subtilisin to generate amino acids 1-20 (RNase S-peptide) and amino acids 21-124 (RNase S-protein). The dark lines between pairs of cysteines represent disulfide bonds.

and their putative KFERQ-like sequences are also listed in Table 6.2. The consensus motif has proven to be correct with some minor but important adjustments; N may substitute for the closely related Q. We have made an RNase S-peptide-β−galactosidase fusion protein and have studied its degradation after introduction into living fibroblasts. The RNase S-peptide causes the β-galactosidase to become a substrate for this pathway of proteolysis, and we have begun examining mutants in the KFERQ region to experimentally verify which peptides can serve as a targeting signal. Our initial results show that QREFK works as well as KFERQ and that P in any position inactivates the signal. (A. Majeski, A. M. Cuervo, L. Terlecky, M. KirvenBrooks, E. Frutiger, and J. F. Dice, unpublished results). An analysis of the conformations of some KFERQ motif peptides based on X-ray crystal structures indicated that they are near the ends of surface helices with one or more side chains buried within the structure.39 The authors concluded that such peptide sequences were unlikely to be recognizable by the intracellular machinery required for this selective pathway of lysosomal proteolysis.39 However, most proteins are much more dynamic than is evident by their x-ray crystal structures. In fact, the KFERQ sequence in RNase A lies entirely within an α-helix, and the F appears to be deeply buried and the E, R, and Q appear to be partially buried.39 Nevertheless, this pentapeptide can be efficiently recognized by antibodies raised against the linear pentapeptide33 and can also be recognized by hsc73.40

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Fig. 6.5. Degradation of RNase A, RNase S-protein, and RNase S-peptide after microinjection into confluent IMR-90 human fibroblasts. Closed circles and solid lines show degradation rates in the presence of serum. Open circles and dashed lines show degradation rates in the absence of serum. The numbers refer to half-lives in hrs.

Reproduction of the Selective Pathway of Lysosomal Proteolysis In Vitro We have been able to reproduce this selective pathway of lysosomal proteolysis using highly purified lysosomes derived from human fibroblasts,35,40,41 Chinese hamster ovary (CHO) cells,49 and rat liver,35,42 kidney,37 and spleen.36 Binding, uptake, and degradation of protein substrates can be separately analyzed in these preparations (Fig. 6.8).43 After


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Fig. 6.6. Identification of amino acids within RNase S-peptide that are required for the increased degradation of microinjected RNase A in response to serum deprivation. The indicated peptides were synthesized, purified, and coloaded into red cell ghosts with radiolabeled RNase A. The ghosts were fused with IMR-90 fibroblasts, and degradation of RNase A in the presence and absence of serum was determined. The ability of the various peptides to inhibit the increased degradation of RNase A in the absence of serum is indicated.

incubation of isolated lysosomes with a substrate protein, most (>90%) of the protein associated with lysosomes is bound to the lysosome surface, because the protein is rapidly degraded once it enters the lysosomal matrix. The amount of substrate transferred into the lysosomal matrix can be quantitated if the lysosomes are incubated in the presence of protease inhibitors and the membrane-bound fraction is removed by treatment with an exogenous protease after the incubation.43 Degradation can be followed using radioactive substrate proteins and following the conversion of acid-precipitable radioactivity to acidsoluble radioactivity.35,41,43 Binding of substrate proteins to the lysosomal membrane is saturable and temperaturedependent.35,41 Substrate proteins compete with each other for binding and uptake.35,36 Lysosomes isolated from cells that have been previously serum-deprived41 or from liver of rats that have been starved42 show an increased ability to bind, take up, and degrade substrate proteins. The binding of substrate proteins increases when a molecular chaperone, hsc73, is added to the purified lysosomes.40-42 This heat shock 70 kilodalton (hsp70) family member also stimulates the import of proteins into the endoplasmic reticulum, mitochondria, and peroxisomes.44-46 HscT3 also stimulates the import of some proteins, but not others, into the nucleus.47 ATP/MgCl2 also stimulates the degradation of substrate proteins, but whether or not the nucleotide stimulates binding and uptake of substrate proteins is not yet known. It is possible that the ATP requirement is only for the ATP-dependent proton pump required to maintain the acidic pH within lysosomes.48

The Receptor in the Lysosomal Membrane The saturable binding of substrate proteins to the lysosomal membrane could be partially inhibited by treating the lysosomes with a previous mild protease,35,41,42 so we suspected the binding was to a lysosomal membrane protein. We separated lysosomal membrane proteins by sodium dodecylsulfate polyacrylamide gel electrophoresis

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Fig. 6.7. Effects of covalent attachment of RNase S-peptide on the degradation of three microinjected proteins. The upper panel shows half-lives of microinjected [3H]RNase S-protein, [3H]lysozyme, and [3H]insulin A chain. Degradation of these proteins is unaffected by serum withdrawal. The bottom panel shows that after covalent attachment of an average of one RNase S-peptide molecule per molecule of [3H]protein, degradation rates of all three proteins are increased by serum withdrawal.

(SDS-PAGE) and then transferred the proteins to nitrocellulose paper.49 When we added RNase A, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), or other substrates for this lysosomal degradation pathway to these preparations, they associated with a protein of 96 kilodaltons (kDa). A variety of proteins that are not substrates show no binding to this 96 kDa lysosomal membrane protein. Using GAPDH-agarose affinity columns, we purified this 96 kDa protein from solubilized lysosomal membranes. Its amino acid sequence


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Table 6.2. Peptide sequences related to KFERQ found in other substrates for the selective lysosomal proteolytic pathway Protein

Sequence

Representation

RNase A

KFERQ

+[]-+Q

Aspartate aminotransferase

RKVEQ

++[]-Q

Pyruvate kinase

QDLKF

Q-[]+[]

Hemoglobin

QRFFE

Q + [ ][ ] -

Aldolase

QKKEL QFREL IKLDQ

Q + + -[ ] Q [ ] + -[ ] [ ] + [ ]- Q

GAPDH

NRVVD

N + [ ][ ]-

IkB

VKELQ

[]+-[]Q

Hsc73

QRDKV QKILD

Q+-+[] Q+[][]-

α2-microglobulin

VDKLN RIKEN

[]-+[]N

Annexin II

QKVFD

Q+[][]-

Annexin IV

QELRR

Q-[]++

Consensus: (+,-,[ ], +/[ ])Q or Q(+,-,[ ],+/[ ]) + = K,R; - = D,E; [ ] = F,I,L,V Nonsubstrates that contain no KFERQ motif include RNase S-protein, lysozyme, ovalbumin, insulin A chain, ubiquitin, β-galactosidase (from E. coli), annexin V, and annexin XI.

was identical to a previously described rat liver lysosomal membrane protein, lysosomal glycoprotein of 96 kDa (LGP96) also referred to as lysosomal-associated membrane protein-2a (LAMP-2a).49 This protein had been studied with regard to how it is targeted to lysosomes (see Chapter 1),50,51 but it did not have a known function. A related protein, LGP120 or LAMP-1, showed no binding to substrate proteins using the same binding assays.49 The structure of LGP96 is shown diagramatically in Fig. 6.9A. We reasoned that the binding of substrate proteins to LGP96 must be through its short 12 amino acid cytosolic tail. This proved to be true since binding could be blocked with excess free peptide with the LGP96 sequence but not with excess peptide with the same amino acid composition, but in a randomized sequence.49 In addition, substrate binding was blocked by an antibody to the cytosolic tail peptide but not by other antibodies.49 More recent results indicate that LGP96 exists as a multimer in the lysosomal membrane (Fig. 6.9B). We obtained the cDNA coding for human LAMP-2a and transfected-CHO cells. We isolated stable transfectants that proved to overexpress LAMP-2a from 26-fold. We then radiolabeled these different cells to follow protein degradation in the presence and absence

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Fig. 6.8. Different steps in selective lysosomal proteolysis. When isolated lysosomes are incubated with a substrate protein and lysosomes are reisolated, most (>85%) of the recovered protein is bound to the lysosomal surface. The substrate is rapidly taken up by lysosomes, but is quickly degraded once in the lysosomal matrix. To measure uptake of the substrate into the lysosomal matrix, the lysosomes are incubated with substrate protein in the presence of chymostatin to inhibit lysosomal cathepsins. After the incubation, proteinase K is added to digest any substrate still bound to the lysosomal surface. The entire process of binding, uptake, and degradation of substrate proteins can be measured as the conversion of acid-precipitable radioactivity to acid-soluble radioactivity.

of serum. The selective lysosomal pathway of proteolysis was increased in direct proportion to the degree of overexpression of LAMP-2a (Fig. 6.10). In addition, this pathway of proteolysis was also activated to some extent even in cells maintained in the presence of serum (Fig. 6.10). These results proved that the level of LGP96 or LAMP-2a was a ratelimiting component for this pathway of proteolysis. Our subsequent studies have shown that levels of lysosomal membrane LGP96 are modulated under physiological and pathological conditions that alter the activity of this selective pathway of lysosomal proteolysis. For example, the activity of liver lysosomes increases progressively during starvation for 3 days,42 and it also increases in kidney lysosomes in response to hydrocarbon exposure.37 In these cases levels of LGP96 in the lysosomal membrane also increase. Lysosomes derived from livers of old rats are less active


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Fig. 6.9. Diagrammatic representation of the receptor in the lysosomal membrane. LGP96 (also known as LAMP2a) has a single transmembrane spanning region with most of the protein within the lysosomal matrix. This domain is highly glycosylated (Y). The carboxyl terminal 12 amino acids that face the cytosol are shown in single letter amino acid abbreviation as defined in the legend to Figure 6.4. It is this cytosolic tail region that specifically interacts with substrate proteins, and this interaction is facilitated by hsc73. Also shown is a tetramer of LGP96 molecules in the lysosomal membrane. Recent work (A. M. Cuervo and J. F. Dice, unpublished results) shows that LGP96 forms tetramers and higher molecular weight structures.

than those from young rats in this selective lysosomal proteolytic pathway, and the lysosomal membrane of old rats contains reduced amounts of LGP96 (A. M. Cuervo and J. F. Dice, unpublished results).

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Fig. 6.10. Effect of overexpression of LGP96 on the selective lysosomal pathway of proteolysis. Chinese hamster ovary cells were stably transfected with a cDNA encoding human LGP96 (hLGP96). The level of LGP96 expression in the colony analyzed was 3-fold above controls. Cellular proteins were radiolabeled with [3H]leucine, and degradation of proteins was followed after switching cell cultures to nonradioactive medium. The LGP96 overexpressing cells showed higher rates of protein degradation both in the presence and in the absence of serum.

Possible Roles of Other Lysosomal Membrane and Matrix Proteins Other lysosomal membrane components may also play a role in the binding and/or uptake of substrate proteins. For example, a portion of hsc73 associates with the lysosomal membrane and may play a role in the passage of substrate proteins through the lysosomal membrane.52 Also, another lysosomal protein can be found associated with the lysosomal matrix side of LGP96. We have identified this protein as cathepsin A (A. M. Cuervo and J.F. Dice, unpublished results). This cathepsin protects a variety of lysosomal enzymes from proteolytic attack in a manner that does not depend on its own proteolytic activity.53,54 Whether or not cathepsin A plays a direct role in this selective pathway of lysosomal proteolysis remains to be established.


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Another critical component in this pathway of proteolysis is a molecular chaperone within the lysosomal lumen.55,56 Hsp70s within the lumen of the ER57,58 and in the matrix of mitochondria4,59 are known to be required to pull the substrate proteins across the membrane. We think the lysosomal hsp70 is actually a population of cytosolic hsc73 that is very stable within lysosomes.55 The lysosomal hsc73 (ly-hsc73) reacts with two different antibodies that are specific for cytosolic hsc73.55 The ly-hsc73 in the lumen is identical in isoelectric point to the most acidic of four isoforms of cytosolic hsc73. Whether or not only this isoform enters lysosomes remains to be established. It is also possible that all forms enter lysosomes but that the three more basic forms are then modified slightly in the lysosomal matrix. We were able to deliver a monoclonal antibody to lysosomes by endocytosis and could inactivate the lysosomal hsc73.55 After this treatment the serum-induced stimulation in protein degradation was completely blocked. Under these conditions, an endocytosed protein was degraded within lysosomes at the normal rate. Therefore, the ly-hsc73 was not affecting the rate of degradation of proteins within lysosomes, so it seemed to be required for the entry of proteins into lysosomes.55 These results were confirmed and expanded by studies of rat liver lysosomes.56 Lysosomes could be separated by slight density differences into two populations. These populations were identical with respect to cathepsin levels in the matrix as well as total LGP96 levels. However, one population of lysosomes contained very little ly-hsc73 in the matrix. This population was relatively inactive in the ability to selectively take up substrate proteins when compared to the population that contained ly-hsc73 in the matrix. However, when this population was preincubated with cytosolic hsc73 so that some hsc73 entered the lysosomal matrix, activity of this population was completely restored.56 An interesting observation in the fibroblasts is that the punctate-appearing lysosomes fuse to form a tubule network when the proteolytic pathway is activated during serum withdrawal.55 This massive fusion may or may not affect lysosomal activities. One possibility is that the fusion corrects for heterogeneity in the lysosomal populations so that, for example, all the lysosomes contain hsc73 as well as LGP96.

Regulation of the Selective Lysosomal Proteolytic Pathway This pathway of proteolysis is activated due to prolonged starvation in rat liver (Fig. 6.11) and due to withdrawal of serum growth factors from confluent cells in culture. The cells in culture have previously induced macroautophagy as a result of reaching confluence, and the rat liver has previously induced macroautophagy due to short term starvation. The rate-limiting step in this selective pathway of proteolysis seems to be the level of LGP96 in the lysosomal membrane. In addition, the amount of hsc73 in the lysosomal matrix can determine the activity under certain conditions. How the levels of hsc73 or LGP96 in lysosomes are regulated is completely unknown. Likewise, the intracellular second messenger systems responsible for increasing the activity of this selective pathway of lysosomal proteolysis are not known with certainty. Recently we obtained evidence implicating oxygen radicals or other oxidizing agents in causing the increased degradation of at least one substrate protein following serum withdrawal.36 Whether or not the oxidation was affecting this specific protein only or also modulates the activity of the selective pathway of lysosomal proteolysis remains to be studied.

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Fig. 6.11. Comparison of timing of activation of macroautophagy and the selective pathway of lysosomal proteolysis in response to fasting. Macroautophagy is stimulated early in starvation and almost returns to normal before the selective pathway is activated. Dotted line = macroautophagy (arbitrary units). Solid line = the selective pathway (arbitrary units).

A Working Model for the Selective Lysosomal Degradation Pathway Our current understanding of this pathway of proteolysis is presented in Fig. 6.12. The targeting sequence related to KFERQ together with additional flanking amino acids is recognized by hsc73. The intracellular mechanisms that regulate this interaction are largely unknown. After the substrate protein has interacted with hsc73, it is better able to bind to LGP96 in the lysosomal membrane. After such binding the substrate protein is internalized through the lysosomal membrane presumably through a gated protein transport channel that has not yet been identified. At least certain protein substrates enter the lysosome by their amino terminus.34 The force for driving the protein to enter the lysosomal matrix appears to be the ly-hsc73. Once in the lysosomal matrix hsc73 detaches from the substrate which is now completely degraded by lysosomal proteases.


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Fig. 6.12. Model for the selective lysosomal pathway of protein degradation. The main steps in the pathway are indicated. 1. Recognition of the substrate protein by cytosolic hsc73, 2. binding of the substrate protein to LGP96, 3. uptake of the substrate protein into the lysosomal matrix with the help of ly-hsc73, 4. degradation of the substrate by lysosomal proteases.

Future Directions of Research The exact role of hsc73 in increasing the binding of substrate proteins to LGP-96 is not known. In particular it will be important to determine whether a substratehsc73 complex binds to LGP96 or if the substrate protein is modified by its interaction with hsc73. Hsp70s usually function in concert with other molecular chaperones called DnaJ homologs.60 We recently found two DnaJ homologs associated with hsc73 within the lysosomal membrane and also in the lysosomal matrix (F. Agarraberes and J. F. Dice, unpublished results). This finding suggests that these forms of ly-hsc73 are functional. We have preliminary evidence to indicate that LGP96 is a multimer in the lysosomal membrane (Fig. 6.9B). This multimerization may be required for conformational changes in the cytosolic tail that are necessary for the tail to bind substrate proteins. The multimerization also suggests the possibility that the transmembrane regions of LGP96 may itself form the aqueous channel through which the substrate proteins travel. Such channels are formed by proteins separate from the receptors in the case of ER4,61 and mitochondrial protein import pathways,4,62 so both functions being carried out by the same protein would be an interesting variation on a theme. If LGP96 is both the receptor and the channel, it should be possible to reconstitute the import using purified LGP96 and artificial liposomes. We could then directly address questions such as lipid and ion requirements and roles for DnaJ homologs for the binding and import. The cytosolic hsc73 binds to a region on the substrate protein that includes the KFERQ peptide motif. However, LGP96 does not seem to bind to this same region of substrate proteins (A.M. Cuervo and J.F. Dice, unpublished results), so mapping out regions of substrate proteins that bind to LGP96 may lead to additional clues about peptide motifs required within substrate proteins to enter this pathway of protein degradation.

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CHAPTER 7

Concluding Remarks Another Pathway for the Delivery of Cytosolic Proteins to the Yeast Vacuole

L

ysosomes are highly dynamic organelles that contain molecular chaperones and several proteases, peptidases, and other hydrolases. There appear to be multiple pathways for the targeting of lysosomal enzymes to the lysosome. This is most evident for the yeast vacuole where genetic analysis combined with biochemical studies has proven that multiple lysosomal targeting pathways exist. Another pathway of internalization of cytosolic proteins by the yeast vacuole has been identified, and mutations in this process are called vacuolar import defective (vid).1-5 These mutations were identified as being defective in the degradation of fructose-1,6-bisphosphatase (FBPase) in yeast that have been refed glucose. These mutations are different from the apg mutants and the ctv mutants described earlier (H.-L. Chiang, personal communication). This pathway involves the selective transport of cytosolic FBPase into novel intracellular vesicles and subsequent fusion of these vesicles with the vacuole.5 This pathway may be unique to yeast. Alternatively, it may represent a variation of the selective lysosomal proteolytic pathway with targeting into a vesicle rather than directly into the lysosome/ vacuole. If this homology is correct, the targeting of FBPase into the intermediate vesicle should be stimulated by hsc73, the intermediate vesicle membrane should contain a receptor, and the lumen of the intermediate vesicles should contain a molecular chaperone to facilitate import. Recent results indicate that the import of FBPase into the intermediate vesicles requires ATP and cytosol (hsc73?) and is saturable (receptor?).6,7 Why yeast would first target proteins to an intermediate vesicle rather than directly to the vacuole is not known. One possibility is that the roles of the yeast vacuole in nonlysosomal functions such as osmotic balance and nutrient storage might be compromised by a gated protein import channel in the vacuolar membrane. Transporting protein substrates first into vesicles may be a simple solution to maintaining a completely sealed vacuolar membrane. Alternatively, the molecular chaperone in the vacuole of yeast may be susceptible to degradation, while hsc73 is resistant to lysosomal degradation in mammalian cells.8 Transporting the protein into an intermediate vesicle that does not contain proteases would bypass this potential problem.


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Index A

I

A-mannosidase 3, 12, 13 Albumin 46, 53, 89 Amino acid 3, 6, 9-12, 21, 22, 27, 32, 39, 45, 46, 57, 59-61, 63-67, 69, 70, 75, 82, 9092, 94, 95, 97, 100 Aminopeptidase I 12 AP-1 8, 45, 46, 48 AP-2 49 ARF 8, 43 Autophagic vacuole 3-5, 13, 57-59, 61-65, 70, 85 Autophagosome 3, 4, 57, 59-63, 65-67, 69-71, 74

Insulin 9, 18, 21, 30, 46, 47, 51-53, 75, 82, 85, 86, 88, 94, 95

C Chaperone 5, 12, 43, 75, 85, 90, 93, 99, 101, 105 Collagen 53 Constitutive secretion 41, 43-45, 51, 54 COPI 43, 44 COPII 8, 43, 44 Cycloheximide 65, 80 Cytoplasm-to-vacuole 69 Cytoskeleton 25, 29, 31, 59, 66, 67, 79

D Dileucine motif 10, 11

F

K KFERQ 90, 91, 94, 100, 101

L LAMP 10, 11, 14, 95-97 LGP96 3, 95-101 LIMP 10, 11, 14

M Mannose-6-phosphate 7, 44, 46 Mannose-6-phosphate receptor 21, 25, 32, 44-46, 48 Methylotrophic yeast 65, 69, 77 Mitochondria 1-6, 57, 69, 85, 93, 99, 101 Multivesicular body 77

P Parathyroid hormone 51, 52 Peroxisome 1-3, 75, 77-82, 85, 93 Peroxisomes 57, 69 PI3 kinase 58, 60, 66, 67 PMSF 80 Prolactin 52 Protein phosphorylation 67 Proton pump 5, 7, 14, 57, 58, 62, 93

Fluid-phase endocytosis 18

G GERL 1 Glucagon 18, 57, 65, 66, 69 Green fluorescent protein 78, 80, 81 GTP-g-S 61, 67, 68

H HSC73 5 Hsc73 26, 48, 90, 91, 93, 94, 97-101, 105

R rab 28, 70 Raffinose 88, 89 Receptor 3, 7, 9, 10, 18-22, 25, 26, 28-30, 32-35, 43-46, 48, 53, 63, 66, 70, 85, 86, 97, 101, 105 Receptor-mediated endocytosis 7, 8, 18, 19, 21, 25, 26, 28, 29, 32, 35 Red cell-mediated microinjection 89 Regulated secretion 39, 43, 48, 50, 51 RNase A 88-92, 94 RNase S-peptide 90-92, 95 RNase S-protein 90-92, 94, 95


108

S

T

S6 58, 67, 69 Sphingomyelinase 29, 79, 82, 83

3-methyladenine 60

U Ubiquitin 32, 34, 62, 63, 69, 70, 94

Lysosomal Pathways of Protein Degradation (Molecular Biology Intelligence Unit)

Lysosomal Pathways of Protein Degradation

Protein Kinase C (Molecular Biology Intelligence Unit)

Endosomes (Molecular Biology Intelligence Unit)

Calreticulin (Molecular Biology Intelligence Unit)

p53 (Molecular Biology Intelligence Unit)

Protein Movement Across Membranes (Molecular Biology Intelligence Unit)

Molecular Biology of the Parathyroid (Molecular Biology Intelligence Unit)

Immunobiology of Carbohydrates (Molecular Biology Intelligence Unit)

Ubiquitin and Disease (Molecular Biology Intelligence Unit)

Translation Mechanisms (Molecular Biology Intelligence Unit)

Cell-Cell Channels (Molecular Biology Intelligence Unit)

Lipocalins (Molecular Biology Intelligence Unit (Unnumbered).)

HOX Gene Expression (Molecular Biology Intelligence Unit)

Rho GTPases (Molecular Biology Intelligence Unit)

Tight Junctions (Molecular Biology Intelligence Unit)

VEGF in Development (Molecular Biology Intelligence Unit)

Branching Morphogenesis (Molecular Biology Intelligence Unit)

Ceramide Signaling (Molecular Biology Intelligence Unit)

The Nucleolus (Molecular Biology Intelligence Unit)

HOX Gene Expression (Molecular Biology Intelligence Unit)

Molecular Basis Of Epithelial Appendage Morphogenesis (Molecular Biology Intelligence Unit)

Molecular Mechanisms of Smooth Muscle Contraction (Molecular Biology Intelligence Unit)

Discovering Biomolecular Mechanisms with Computational Biology (Molecular Biology Intelligence Unit)

Discovering Biomolecular Mechanisms with Computational Biology (Molecular Biology Intelligence Unit)

The Biogenesis of Cellular Organelles (Molecular Biology Intelligence Unit)

Regulation of G1 Phase Progression (Molecular Biology Intelligence Unit)

Proteasomes : The World of Regulatory Proteolysis (Molecular Biology Intelligence Unit)

Molecular Mechanisms of Phagocytosis (Medical Intelligence Unit)

Trafficking Inside Cells: Pathways, Mechanisms and Regulation (Molecular Biology Intelligence Unit)

Voltage-Gated Calcium Channels (Molecular Biology Intelligence Unit)

Lysosomal Pathways of Protein Degradation (Molecular Biology Intelligence Unit)