Methods in
Molecular Biology 1594

Karin Öllinger
Hanna Appelqvist Editors

Lysosomes
Methods and Protocols


Methods

in

Molecular Biology

Series Editor
John M. Walker
School of Life and Medical Sciences
University of Hertfordshire
Hatfield, Hertfordshire, AL10 9AB, UK

For further volumes:
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Lysosomes
Methods and Protocols

Edited by

Karin Öllinger

Experimental Pathology, Department of Clinical and Experimental Medicine, Linköping University,
Linköping, Sweden

Hanna Appelqvist
Department of Physics, Chemistry and Biology, Linköping University, Linköping, Sweden


Editors
Karin Öllinger
Experimental Pathology
Department of Clinical and Experimental
Medicine
Linköping University
Linköping, Sweden

Hanna Appelqvist
Department of Physics, Chemistry and Biology
Linköping University
Linköping, Sweden

ISSN 1064-3745    ISSN 1940-6029 (electronic)
Methods in Molecular Biology
ISBN 978-1-4939-6932-6    ISBN 978-1-4939-6934-0 (eBook)
DOI 10.1007/978-1-4939-6934-0
Library of Congress Control Number: 2017935483
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Preface
The endo-lysosomal system is central to the degradation and recycling of macromolecules
delivered by endocytosis, phagocytosis, and autophagy [1–3]. As the major digestive compartment within cells, lysosomes harbor around 60 acidic hydrolases, responsible for the
cellular digestion of most macromolecules. The lysosomal function goes far beyond the
degradation activity and lysosomes are identified as important regulators of nutrient sensing, exocytosis, receptor recycling and regulation, cell death, and cholesterol homeostasis
[4–7]. A significant finding recognized lysosomes as important signaling organelles that
sense nutrient availability and generate an adaptive response to maintain cellular homeostasis, mainly through activation of the transcription factor EB (TFEB) [8]. The discovery of
TFEB as a master regulator of lysosomal biogenesis, regulator of autophagic function and
energy metabolism has greatly impacted our view of lysosomes as important hubs for interpretation of environmental alterations [9]. In addition, the lysosomes function as a Ca2+
store that participates in the signal transduction eventually leading to the nuclear translocation of TFEB [10]. The importance of lysosomes for cellular cholesterol homeostasis was
identified through the inherited lysosomal storage disorder Niemann-Picks disease type C,
which is caused by mutation in either of the two proteins NPC1 and NPC2 [11].
Furthermore, a lysosomal hydrolase-mediated digestion of LDL and subsequent cholesterol release from the lysosomes through the action of NPC1 and NPC2, by a not yet fully
defined mechanism, has also recognized the importance of lysosomes in atherosclerosis
[12].
Moreover, the lysosome is centrally involved in the regulation and control of cell death
and survival. Due to their high content of hydrolytic enzymes, lysosomes are potentially

harmful to cells. Christian de Duve termed the lysosomes “suicide bags” as massive lysosomal rupture may cause cytosolic acidification followed by necrosis [13]. Present knowledge has however shown that partial and selective lysosomal membrane permeabilization
(LMP) could trigger several forms of controlled cell death [14]. LMP results in the release
of lysosomal content to the cytosol and the main lysosomal hydrolases implicated in triggering of cell death are the cathepsins, which have been shown in several in vitro system but also
in vivo [5, 15–17]. The mechanism of LMP is not clarified and most likely lysosomal permeabilization is due to alteration in both lysosomal membrane proteins and lipids causing
destabilization of the membrane. Interestingly, in addition to the role of lysosomes in cell
death they are also involved in the repair of the plasma membrane. In response to plasma
membrane rupture, lysosomes are able to rescue the cell by rapid translocation to the damage site of the plasma membrane and donation of the membrane [18, 19]. This exocytosis
process is triggered by Ca2+ influx from the extracellular compartment and requires the
ubiquitously expressed lysosomal membrane protein synaptotagmin 7 [20]. Besides conventional lysosomes, lysosome-related organelles (LRO), including melanosomes, lytic granules, and platelet-dense granules, exist in certain cell types and have acquired special functions
[21].
Over the last decade, advances in lysosome research have established a broad role for
the lysosomes in the pathophysiology of disease. The most obvious are the lysosomal stor-

v


vi

Preface

age diseases (LSD), which include approximately 70 distinct disorders. Although individually rare, they collectively account for 14 % of all inherited metabolic diseases. The main
biochemical hallmark of LSD is the accumulation of un- or partially digested metabolites in
the lysosomes. The pathologic mechanisms include malfunction of the degradation, the
transport across the lysosomal membrane, or trafficking between endosomes and lysosomes
[22]. Noteworthy, recent studies have observed that lysosomal alterations and malfunction
are also players in some of the most common conditions nowadays including cancer and
neurodegenerative diseases. The neurodegenerative hallmarks of the rare early-­onset lysosomal storage diseases resemble late-onset neurodegenerative diseases such as Alzheimer’s
and Parkinson’s diseases. It has been shown that type 1 Gaucher disease patients have a
higher risk of developing Parkinson’s disease [23]. Frontotemporal dementia is caused by
mutation in one allele of progranulin. However if both alleles are mutated, it will lead to

the neuronal ceroid lipofuscinogenesis (CLN11) [24]. Thus a theory of a general mechanism of dysfunctional clearance of cellular cargo through the secretory-endosomal-­
autophagic-lysosomal-exocytic (SEALE) network has been formed to explain the common
underlying feature relating lysosomal dysfunction to seemingly different diseases [25].
Advanced tumor cells are highly dependent on effective lysosomal function. Thus, cancer progression and metastasis are associated with striking alterations in lysosomal compartments including changes in lysosome volume, composition, cellular distribution, and
lysosomal enzyme activity. Release of cathepsins from a cancer cell into the extracellular
space can promote tumor growth through their proteolytic effect on the basement membrane and activation of other pro-tumorigenic proteins [26–28]. Moreover, elevated
expression of wild-type TFEB protein is sufficient for driving the oncogenic mechanism
[29]. Resistance of cancer cells towards traditional therapies may be overcome by agents
that trigger LMP and engage lysosomal cell death pathways [26]. On the other hand,
therapeutic strategies to restrain proteolytic activity of secreted hydrolases would be a way
to suppress tumor invasion. The development of techniques for control and manipulation
of lysosomal function will generate future treatments of the wide variety of common and
rare pathological conditions involving lysosomes.
After several groundbreaking discoveries, our knowledge has increased tremendously
and the lysosome is now recognized as one of the central organelles for normal physiological function and during disease. In this volume of Methods in Molecular Biology, laboratory protocols for detailed studies of essential parts of lysosomal biology are provided. The
protocols are straightforward and aim to guide researchers in their exploration of lysosomes, both under normal conditions and in pathological processes. We hope that the
provided know-how and protocols will guide and inspire further research and generate new
insights into the versatile tasks of this fascinating organelle.
Finally, we would like to thank all contributing authors for sharing their expertise. We
would also express our sincere gratitude to Professor John M. Walker for support and guidance during the editing of this volume of MiMB series.
Linköping, Sweden
Linköping, Sweden

Karin Öllinger
Hanna Appelqvist


Preface

vii


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24.Smith KR, Damiano J, Franceschetti S, Carpenter S, Canafoglia L, Morbin M, Rossi G, Pareyson D,
Mole SE, Staropoli JF, Sims KB, Lewis J, Lin WL, Dickson DW, Dahl HH, Bahlo M, Berkovic SF
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Contents
Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
v
Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xi
  1 SILAC-Based Comparative Proteomic Analysis of Lysosomes
from Mammalian Cells Using LC-MS/MS . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Melanie Thelen, Dominic Winter, Thomas Braulke,
and Volkmar Gieselmann
  2 Quantitative Profiling of Lysosomal Lipidome by Shotgun Lipidomics . . . . . . .
Mesut Bilgin, Jesper Nylandsted, Marja Jäättelä, and Kenji Maeda
  3 Analysis of N- and O-Glycosylation of Lysosomal Glycoproteins . . . . . . . . . . . .
Elmira Tokhtaeva, Olga A. Mareninova, Anna S. Gukovskaya,
and Olga Vagin
  4 Analyzing Lysosome-Related Organelles by Electron Microscopy . . . . . . . . . . .
Ilse Hurbain, Maryse Romao, Ptissam Bergam, Xavier Heiligenstein,
and Graça Raposo
  5 Microscopic Analysis of Lysosomal Membrane Permeabilization . . . . . . . . . . . .
Ana Maria Vilamill Giraldo, Karin Öllinger, and Vesa Loitto
  6 Quantitative Co-Localization and Pattern Analysis of Endo-­L ysosomal

Cargo in Subcellular Image Cytometry and Validation on Synthetic
Image Sets . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Frederik W. Lund and Daniel Wüstner
  7 Preparation of a Two-Photon Fluorescent Probe for Imaging H2O2
in Lysosomes in Living Cells and Tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Mingguang Ren, Beibei Deng, Xiuqi Kong, Yonghe Tang,
and Weiying Lin
  8 Lysophagy: A Method for Monitoring Lysosomal Rupture Followed
by Autophagy-Dependent Recovery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Takanobu Otomo and Tamotsu Yoshimori
  9 Delivery of Cargo to Lysosomes Using GNeosomes . . . . . . . . . . . . . . . . . . . . .
Kristina M. Hamill, Ezequiel Wexselblatt, Wenyong Tong,
Jeffrey D. Esko, and Yitzhak Tor
10 Lysosomal Acidification in Cultured Astrocytes Using Nanoparticles . . . . . . . .
Camilla Lööv and Anna Erlandsson
11 Analysis of Lysosomal pH by Flow Cytometry Using FITC-­Dextran
Loaded Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Ida Eriksson, Karin Öllinger, and Hanna Appelqvist
12 Detection of Lysosomal Exocytosis in Platelets by Flow Cytometry . . . . . . . . .
Anna L. Södergren and Sofia Ramström

ix

1

19
35

43




73

93

129

141
151

165

179
191


x

Contents

13 Detection of Lysosomal Exocytosis by Surface Exposure of Lamp1
Luminal Epitopes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Norma W. Andrews
14 Using the MEROPS Database for Investigation of Lysosomal Peptidases,
Their Inhibitors, and Substrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Neil D. Rawlings
15 Next-Generation Sequencing Approaches to Define the Role
of the Autophagy Lysosomal Pathway in Human Disease:
The Example of LysoPlex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Giuseppina Di Fruscio, Sandro Banfi, Vincenzo Nigro,
and Andrea Ballabio
16 Gelatin Zymography Using Leupeptin for the Detection
of Various Cathepsin L Forms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Yoko Hashimoto
17 Methods for Determination of α-Glycosidase, β-Glycosidase,
and α-Galactosidase Activities in Dried Blood Spot Samples . . . . . . . . . . . . . . .
Eser Yıldırım Sozmen and Ebru Demirel Sezer
18 Prenatal Diagnosis of Lysosomal Storage Disorders Using Chorionic Villi . . . .
Jyotsna Verma, Sunita Bijarnia-Mahay, and Ishwar C. Verma
19 Lysosomal Biology in Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Colin Fennelly and Ravi K. Amaravadi

205

213

227

243

255
265
293

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 309


Contributors
Ravi K. Amaravadi  •  Department of Medicine and Abramson Cancer Center, Perelman

School of Medicine, University of Pennsylvania, Philadelphia, PA, USA
Norma W. Andrews  •  Department of Cell Biology and Molecular Genetics, University
of Maryland at College Park, College Park, MD, USA
Hanna Appelqvist  •  Division of Chemistry, Department of Physics, Chemistry and Biology,
Linköping University, Linköping, Sweden
Andrea Ballabio  •  Telethon Institute of Genetics and Medicine (TIGEM), Pozzuoli (NA),
Italy; Medical Genetics, Department of Translational Medicine, Federico II University,
Naples, Italy; Department of Molecular and Human Genetics, Baylor College of
Medicine, Houston, TX, USA; Jan and Dan Duncan Neurological Research Institute,
Texas Children’s Hospital, Houston, TX, USA
Sandro Banfi  •  Medical Genetics, Department of Biochemistry, Biophysics and General
Pathology, Second University of Naples, Naples, Italy; Telethon Institute of Genetics and
Medicine (TIGEM), Pozzuoli (NA), Italy
Ptissam Bergam  •  Institut Curie, PSL Research University, CNRS, Paris, France;
Sorbonne Universités, UPMC Univ Paris 06, CNRS, Paris, France; Cell and Tissue
Imaging Core Facility PICT-IBiSA, Institut Curie, Paris, France; Characterization
Core Lab, King Abdullah University of Science and Technology (KAUST), Thuwal,
Kingdom of Saudi Arabia
Sunita Bijarnia-Mahay  •  Institute of Medical Genetics and Genomics, Sir Ganga Ram
Hospital, New Delhi, India
Mesut Bilgin  •  Cell Death and Metabolism Unit, Center for Autophagy, Recycling
and Disease, Danish Cancer Society Research Center, Copenhagen, Denmark
Thomas Braulke  •  Department of Biochemistry, Children’s Hospital, University Medical
Center Hamburg-Eppendorf, Hamburg, Germany
Beibei Deng  •  Institute of Fluorescent Probes for Biological Imaging, School of Chemistry
and Chemical Engineering, School of Materials Science and Engineering, University of
Jinan, Jinan, Shandong, P.R. China
Giuseppina Di Fruscio  •  Medical Genetics, Department of Biochemistry, Biophysics
and General Pathology, Second University of Naples, Naples, Italy
Ida Eriksson  •  Experimental Pathology, Department of Clinical and Experimental

Medicine, Linköping University, Linköping, Sweden
Anna Erlandsson  •  Department of Public Health and Caring Sciences/Molecular
Geriatrics, Rudbeck Laboratory, Uppsala University, Uppsala, Sweden
Jeffrey D. Esko  •  Cellular and Molecular Medicine, University of California, San Diego,
La Jolla, CA, USA
Colin Fennelly  •  Department of Medicine and Abramson Cancer Center, Perelman
School of Medicine, University of Pennsylvania, Philadelphia, PA, USA
Volkmar Gieselmann  •  Institute for Biochemistry and Molecular Biology,
Rheinische-­Friedrich-­Wilhelms-University, Bonn, Germany
Ana Maria Vilamill Giraldo  •  Experimental Pathology, Department of Clinical
and Experimental Medicine, Linköping University, Linköping, Sweden

xi


xii

Contributors

Anna S. Gukovskaya  •  David Geffen School of Medicine, University of California at Los
Angeles, Los Angeles, CA, USA; VA Greater Los Angeles Healthcare System, Los Angeles,
CA, USA
Kristina M. Hamill  •  Department of Chemistry and Biochemistry, University of
California, San Diego, La Jolla, CA, USA
Yoko Hashimoto  •  Department of Biochemistry, School of Dentistry, Aichi-Gakuin
University, Chikusa-ku, Nagoya, Japan
Xavier Heiligenstein  •  Institut Curie, PSL Research University, CNRS, Paris, France;
Sorbonne Universités, UPMC Univ Paris 06, CNRS, Paris, France; Cell and Tissue
Imaging Core Facility PICT-IBiSA, Institut Curie, Paris, France
Ilse Hurbain  •  Institut Curie, PSL Research University, CNRS, Paris, France; Sorbonne

Universités, UPMC Univ Paris 06, CNRS, Paris, France; Cell and Tissue Imaging Core
Facility PICT-IBiSA, Institut Curie, Paris, France
Marja Jäättelä  •  Cell Death and Metabolism Unit, Center for Autophagy, Recycling
and Disease, Danish Cancer Society Research Center, Copenhagen, Denmark
Xiuqi Kong  •  Institute of Fluorescent Probes for Biological Imaging, School of Chemistry
and Chemical Engineering, School of Materials Science and Engineering, University of
Jinan, Jinan, Shandong, P.R. China
Weiying Lin  •  Institute of Fluorescent Probes for Biological Imaging, School of Chemistry
and Chemical Engineering, School of Materials Science and Engineering, University of
Jinan, Jinan, Shandong, P.R. China
Vesa Loitto  •  Core Facility Microscopy Unit, Medical Faculty, Linköping University,
Linköping, Sweden
Camilla Lööv  •  MassGeneral Institute for Neurodegeneration, Massachusetts General
Hospital, Harvard Medical School, Charlestown, MA, USA
Frederik W. Lund  •  Department of Biochemistry and Molecular Biology, University of
Southern Denmark, Odense M, Denmark; Department of Biochemistry, Weill Medical
College of Cornell University, New York, NY, USA
Kenji Maeda  •  Cell Death and Metabolism Unit, Center for Autophagy, Recycling and
Disease, Danish Cancer Society Research Center, Copenhagen, Denmark
Olga A. Mareninova  •  David Geffen School of Medicine, University of California at Los
Angeles, Los Angeles, CA, USA; VA Greater Los Angeles Healthcare System, Los Angeles,
CA, USA
Vincenzo Nigro  •  Medical Genetics, Department of Biochemistry, Biophysics and General
Pathology, Second University of Naples, Naples, Italy; Telethon Institute of Genetics and
Medicine (TIGEM), Pozzuoli (NA), Italy
Jesper Nylandsted  •  Cell Death and Metabolism Unit, Center for Autophagy, Recycling
and Disease, Danish Cancer Society Research Center, Copenhagen, Denmark
Karin Öllinger  •  Experimental Pathology, Department of Clinical and Experimental
Medicine, Linköping University, Linköping, Sweden
Takanobu Otomo  •  Department of Genetics, Osaka University Graduate School of

Medicine, Osaka, Japan; Laboratory of Intracellular Membrane Dynamics, Osaka
University Graduate School of Frontier Biosciences, Osaka, Japan; Research Center for
Autophagy, Osaka University Graduate School of Medicine, Osaka, Japan
Sofia Ramström  •  Department of Clinical Chemistry and Department of Clinical and
Experimental Medicine, Linköping University, Linköping, Sweden; Department of
Clinical Medicine, Örebro University, Örebro, Sweden


Contributors

xiii

Graça Raposo  •  Institut Curie, PSL Research University, CNRS, Paris, France; Sorbonne
Universités, UPMC Univ Paris 06, CNRS, Paris, France; Cell and Tissue Imaging Core
Facility PICT-IBiSA, Institut Curie, Paris, France
Neil D. Rawlings  •  European Molecular Biology Laboratory, European Bioinformatics
Institute (EMBL-EBI), Wellcome Genome Campus, Hinxton, Cambridge, UK
Mingguang Ren  •  Institute of Fluorescent Probes for Biological Imaging, School of
Chemistry and Chemical Engineering, School of Materials Science and Engineering,
University of Jinan, Jinan, Shandong, P.R. China
Maryse Romao  •  Institut Curie, PSL Research University, CNRS, Paris, France; Sorbonne
Universités, UPMC Univ Paris 06, CNRS, Paris, France; Cell and Tissue Imaging Core
Facility PICT-IBiSA, Institut Curie, Paris, France
Ebru Demirel Sezer  •  Department of Medical Biochemistry and Metabolism Laboratory,
Ege University Faculty of Medicine, Izmir, Turkey
Anna L. Södergren  •  Clinical Chemistry, Department of Clinical and Experimental
Medicine, Linköping University, Linköping, Sweden
Eser Yıldırım Sozmen  •  Department of Medical Biochemistry and Metabolism Laboratory,
Ege University Faculty of Medicine, Izmir, Turkey
Yonghe Tang  •  Institute of Fluorescent Probes for Biological Imaging, School of Chemistry

and Chemical Engineering, School of Materials Science and Engineering, University of
Jinan, Jinan, Shandong, P.R. China
Melanie Thelen  •  Institute for Biochemistry and Molecular Biology, Rheinische-­Friedrich-­
Wilhelms-University, Bonn, Germany
Elmira Tokhtaeva  •  David Geffen School of Medicine, University of California at Los
Angeles, Los Angeles, CA, USA; VA Greater Los Angeles Healthcare System, Los Angeles,
CA, USA
Wenyong Tong  •  Cellular and Molecular Medicine, University of California, San Diego,
La Jolla, CA, USA
Yitzhak Tor  •  Department of Chemistry and Biochemistry, University of California, San
Diego, La Jolla, CA, USA
Olga Vagin  •  David Geffen School of Medicine, University of California at Los Angeles, Los
Angeles, CA, USA; VA Greater Los Angeles Healthcare System, Los Angeles, CA, USA
Ishwar C. Verma  •  Institute of Medical Genetics and Genomics, Sir Ganga Ram Hospital,
New Delhi, India
Jyotsna Verma  •  Institute of Medical Genetics and Genomics, Sir Ganga Ram Hospital,
New Delhi, India
Ezequiel Wexselblatt  •  Department of Chemistry and Biochemistry, University of
California, San Diego, La Jolla, CA, USA
Dominic Winter  •  Institute for Biochemistry and Molecular Biology, Rheinische-­Friedrich-­
Wilhelms-University, Bonn, Germany
Daniel Wüstner  •  Department of Biochemistry and Molecular Biology, University
of Southern Denmark, Odense M, Denmark
Tamotsu Yoshimori  •  Department of Genetics, Osaka University Graduate School
of Medicine, Osaka, Japan; Laboratory of Intracellular Membrane Dynamics, Osaka
University Graduate School of Frontier Biosciences, Osaka, Japan; Research Center for
Autophagy, Osaka University Graduate School of Medicine, Osaka, Japan


Chapter 1

SILAC-Based Comparative Proteomic Analysis
of Lysosomes from Mammalian Cells Using LC-MS/MS
Melanie Thelen, Dominic Winter, Thomas Braulke,
and Volkmar Gieselmann
Abstract
Mass spectrometry-based proteomics of lysosomal proteins has led to significant advances in understanding lysosomal function and pathology. The ever-increasing sensitivity and resolution of mass spectrometry
in combination with labeling procedures which allow comparative quantitative proteomics can be applied
to shed more light on the steadily increasing range of lysosomal functions. In addition, investigation of
alterations in lysosomal protein composition in the many lysosomal storage diseases may yield further
insights into the molecular pathology of these disorders. Here, we describe a protocol which allows to
determine quantitative differences in the lysosomal proteome of cells which are genetically and/or biochemically different or have been exposed to certain stimuli. The method is based on stable isotope labeling of amino acids in cell culture (SILAC). Cells are exposed to superparamagnetic iron oxide particles
which are endocytosed and delivered to lysosomes. After homogenization of cells, intact lysosomes are
rapidly enriched by passing the cell homogenates over a magnetic column. Lysosomes are eluted after
withdrawal of the magnetic field and subjected to mass spectrometry.
Key words Lysosome, Magnetic particle, Lysosomal proteome, Lysosomal storage disorder, Mass
spectrometry

1  Introduction
Lysosomes are membrane-limited organelles with an acidic pH
maintained by integral vacuolar H+ ATPases. For a long time, it
was assumed that the sole function of lysosomes is the degradation
of a wide variety of macromolecules and the release of degradation
products into the cytosol, where they can be reused for biosynthetic or energy-producing pathways. During the last years, however, it became clear that lysosomes are not merely degradative
compartments but communicate with their environment and play
important roles in secretion, plasma membrane repair, and antigen presentation [1]. Furthermore, protein complexes localized
at the cytosolic surface of lysosomes, such as mTORC1
Karin Öllinger and Hanna Appelqvist (eds.), Lysosomes: Methods and Protocols, Methods in Molecular Biology, vol. 1594,
DOI 10.1007/978-1-4939-6934-0_1, © Springer Science+Business Media LLC 2017

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Melanie Thelen et al.

(mammalian ­target of rapamycin complex 1) and BORC (biogenesis of lysosome-­related organelles complex 1-related complex),
mediate nutrient signaling and lysosomal positioning, respectively
[2, 3]. This wide spectrum of lysosomal functions requires the
action of many proteins.
Lysosomal degradation of macromolecules depends on about
60 soluble acid hydrolases residing in the lysosomal lumen [4].
The surrounding lysosomal membrane contains numerous integral
membrane proteins encompassing transporters for delivery of degradation products, ion channels or highly glycosylated proteins
securing lysosomal integrity by protecting the lysosomal membrane from self-digestion [5, 6]. Peripheral membrane proteins
and soluble proteins bound to the cytoplasmic side of lysosomes by
protein–protein interactions allow for communication or fusion
with other cellular compartments and integrate the lysosome into
the overall cellular metabolism. Proteomics of soluble lysosomal
proteins revealed important aspects of lysosomal function. In these
studies, lysosomal proteins were enriched using an affinity matrix
that specifically binds the unique mannose 6-phosphate (M6P)
residues found on N-linked oligosaccharide side chains of soluble
proteins in the lysosomal lumen. These investigations provided
novel insights in (1) cell type-specific lysosomal enzymes [7, 8], (2)
binding preferences of the two M6P receptors for subpopulations
of lysosomal enzymes [9, 10], and secreted lysosomal enzymes
found in the circulation [11] or urine [12]. This approach has also
been successfully applied to identify enzyme defects underlying
lysosomal storage disorders such as the cholesterol binding lysosomal protein NPC2 being deficient in a form of Niemann Pick

Type C disease [13]. The success of proteomics of soluble lysosomal hydrolases promises that studies on the comparatively underexplored proteome of the lysosomal membrane, and proteins
associated with the cytosolic surface of lysosomes, will yield new
important insights into lysosomal function. Multi-step subcellular
fractionation techniques for enriching lysosomal membranes and
subsequent mass spectrometric analyses led to the description of
140–300 membrane proteins of variable abundancy with known or
presumed lysosomal localization [5, 14]. Most of these studies
were targeted at the identification of novel bona fide lysosomal
proteins, and have indeed revealed previously unknown proteins of
lysosomal localization. Although the precise function of most of
these proteins is still unknown, their investigation should increase
our understanding of this multifaceted organelle in the future.
Irrespective of the goal of the study, lysosomal proteomics
requires the enrichment of lysosomes which can be achieved by
various techniques. These include subcellular fractionation methods using different density gradient materials like sucrose, percoll,
nycodenz and metrizamide [15, 16], or immunoprecipitation of
lysosomes from LAMP1-FLAG overexpressing cells [17].


Isolation and MS-Analysis of Lysosomes

3

Alternatively, lysosomes can be isolated by using an in vivo approach
employing a density shift of mouse or rat liver lysosomes after injection of Triton WR-1339 [16]. One disadvantage of most density-­
gradient-based approaches, is the contamination with other
organelles, in particular mitochondria. For mass spectrometric
investigations we have adapted a technique using superparamagnetic iron oxide particles for rapid enrichment of lysosomes from
cell homogenates recovering routinely up to 80% of intact lysosomes. For a detailed description of this method see Walker and
Lloyd-Evans [18]. The method relies on the endocytic uptake of

dextran-coated iron oxide particles with 10 nm diameter, which are
subsequently enriched in the lysosomal compartment and can be
isolated by passing the cell homogenate over a magnetic column.
This allows for a rapid enrichment of intact lysosomes in amounts
sufficient for mass spectrometric analysis from as little as two confluent 10 cm dishes of cells. The method is fast, avoids unnecessary
manipulations and is thus likely to preserve the original protein
composition of the sample. Due to the high sensitivity of mass spectrometers, in any proteomic dataset of subcellular fractions, a considerable number of proteins identified cannot be assigned to the
enriched organelles. Some of these proteins are true contaminants
but in case of lysosomes may also co-purify because they are functionally linked to the diverse functions of lysosomes. Lysosomes
have contact sites with the endoplasmic reticulum and mitochondria [19–21], are tightly connected to the cytoskeleton to ensure
their migration within the cytoplasm [22] and through selective
and non-selective autophagy are the ultimate destination of ribosomes, mitochondria, parts of the ER and many cytosolic proteins
[23–25]. Consequently, when applying a highly sensitive analytical
approach, the identification of a wide range of non-lysosomal proteins cannot be avoided and it can therefore be difficult to identify
the proteins which are affected by the experiment. This problem
can be considerably reduced if proteomic datasets of two samples
are quantitatively compared. These samples can differ in genetic
background, be cultured under different conditions or be exposed
to specific pharmacological or biochemical stimuli. In the comparative quantitative proteomic dataset only the amount of those proteins which are somehow functionally connected to the investigated
alteration of the system will change, whereas the numerous irrelevant or contaminating proteins detected in both samples remain
unchanged. Moreover, the nature of the applied stimulus or condition may allow developing a working hypothesis on how the identified proteins are functionally connected to the lysosome. To allow
for quantitative proteomic comparison of lysosomal proteins, we
have used stable isotope labeling of amino acids in cell culture
(SILAC) [26]. Cells with unlabeled amino acids can serve as controls, those labeled with heavy amino acids can differ, e.g. genetically or biochemically, and vice versa.


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Melanie Thelen et al.


We have recently used this technique successfully for the comparative investigation of the proteome of lysosomal hydrolases in
mouse cells lacking M6P targeting signals. This led to the identification of alternative M6P-independent transport pathways and the
receptors involved [27].

2  Materials
2.1  Isolation
of Lysosomes

1.
Human embryonic kidney (HEK) 293 cells (German
Collection of Microorganisms and Cell Cultures, DSMZ).
2. Phosphate buffered saline (PBS).
3. Amino acids l-Lysine-2HCl (13C6, 15N2 labeled and unlabeled,
respectively), 17.45 g/l in PBS (200-fold stock solution) and
l-Arginine-HCL (13C6, 15N4 labeled and unlabeled, respectively) 36.24 g/l in PBS (200-fold stock solution) (see Note 1).
4. Fetal Bovine Serum for SILAC (dialyzed).
5.DMEM for SILAC: Dulbecco’s modified Eagle Medium
(DMEM) high glucose (4.5 g/l) for SILAC, without lysine (Lys)
and arginine (Arg) supplemented with 10% FBS, GlutaMax™
(200 mM), Penicillin (100 U/ml)/streptomycin (0.1 mg/ml)
and either conventional light Arg/Lys or heavy isotope labeled
Arg (13C615N4)/Lys (13C615N2). The final concentrations of Arg
should be 87.8 mg/l and Lys: 181.2 mg/l (see Note 1).
6.Magnetite solution: EndoMAG40, 40 kDa dextran-coated
magnetite particles from Liquids Research ( /> 7. PLL solution: Poly-l-Lysine 0.5 mg/ml in PBS.
8.Isolation buffer: 250 mM sucrose, 10 mM HEPES/OH pH
7.4, 1 mM CaCl2, 1 mM MgCl2, 1.5 mM MgAc, 1 mM dithiothreitol (DTT), 1× protease inhibitor cocktail (PIC, Halt protease inhibitor cocktail). Add DTT and PIC always immediately
before using the buffer.
9. Tight-fitting 7 ml Dounce homogenizer.
10. BSA solution: 0.5 mg/ml bovine serum albumin (BSA) in PBS.

11. DNAse 1 solution: 10 μl DNAse 1 (1 Unit/μl) in 1 ml of isolation buffer.
12. Miltenyi LS Separation columns.
13. Miltenyi MidiMACS Magnetic Separator.

2.2 
β-Hexosaminidase
Enzyme Assay

1.Substrate solution: 10 mM 4-Nitrophenyl N-acetyl-β-Dglucosaminide in 0.1 M sodium citrate, pH 4.6, containing
0.2% BSA.
2. 10% Triton X-100 (v/v) in water.
3. Stop solution: 400 mM glycine/OH pH 10.4.


Isolation and MS-Analysis of Lysosomes

2.3  Mass
Spectrometry Sample
Preparation

5

Prepare all solutions for mass spectrometry sample preparation
with MS grade water and use MS-grade/ultrapure chemicals.
1. Maximum recovery pipet tips.
2. Maximum recovery microcentrifuge tubes.
3. Amicon Ultra Centrifugal Filters Ultracel 3K, Merck Millipore
Ltd.
4.Laemmli buffer 4× concentrated (modified from [28]): 250
mM Tris, 8% (w/v) SDS, 40% (v/v) glycerol, 10% (v/v)

β-mercaptoethanol, 0.004% (w/v) bromophenol blue.
5. 40% (w/v) acrylamide in water.
6. SDS-PAGE Gel (10% acrylamide).
7.PageBlue™ protein staining solution, Fermentas, or any other
Coomassie G250 gel staining solution.
8.Solution A: 30% (v/v) acetonitrile (ACN) in 100 mM
NH4HCO3, pH 7.8.
9. 100 mM NH4HCO3, pH 7.8.
10.Trypsin solution: 0.5 ng/μl sequencing grade trypsin in 50
mM NH4HCO3, pH 7.8. To prepare this solution, resuspend
20 μg of trypsin in 200 μl of water, then mix with 200 μl of
100 mM NH4HCO3, pH 7.8 (see Note 2).
11. Solution B: 0.1% (v/v) trifluoroacetic acid, 50% ACN.
12. Solution C: 5% (v/v) ACN, 5% formic acid (FA).
13. StageTips, prepared with Solid Phase Extraction Disk Octadecyl
C18 (3 M, www.3m.com), 4 layers in a 200 μl Maximum
recovery pipet tip [29].
14. 100% methanol.
15. 0.5% (v/v) acetic acid in 80% ACN.
16. 0.5% (v/v) acetic acid.
17. Vacuum centrifuge.

2.4  LC-MS-MS
Measurement

1.C18 Analytical column: ESI spray tip produced in house with a
Sutter P2000 laser puller device from 360 μm OD, 100 μm ID
fused silica capillary packed with 5 μm particles [Dr. Maisch,
Reprosil C-18 AQ]. Commercially available columns can alternatively be used.
2.Thermo EASY-nLC 1,000 or similar nanoflow high or ultra-­

high performance liquid chromatography systems.
3.Thermo Orbitrap Velos Mass Spectrometer or any other suitable mass spectrometer.
4. Running buffer A: water with 0.1% FA.
5. Running buffer B: ACN with 0.1% FA.


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Melanie Thelen et al.

3  Methods
The general workflow of the described method is graphically displayed in Fig. 1. Perform all steps involving cell culture in a sterile
laminar flow hood. For cell culture, pre-warm all solutions to 37 °C.
3.1  Isolation
of Lysosomes

1.Cultivate the cells for at least six passages (to ensure complete
SILAC labeling of the cells’ proteome) in DMEM for SILAC.
2. The following procedures are described for HEK293 cells. When
you want to use another cell type, please refer to Notes 3–6.
3. For HEK293 cells, 10 cm cell culture dishes need to be coated
in advance with PLL-solution for 10 min at room temperature
(RT) to ensure proper cell attachment. Thereafter, remove the
liquid and wash thrice with PBS before plating the cells
(see Note 7).

Fig. 1 Schematic representation of the experimental workflow. Two cell populations are labeled with either
light or heavy arginine or lysine. They can be exposed to different conditions at any time between seeding and
harvest. After 24 h of incubation with magnetite-containing medium and 24-36 h of chase time, cells are
pooled immediately after harvesting, homogenized, the postnuclear supernatants (PNS) prepared and passed

over a magnetic column. The eluate is then fractionated and proteolytically digested before LC-MS/MS measurement (modified and reproduced from [27] with permission from Wiley)


Isolation and MS-Analysis of Lysosomes

7

4.For each experiment, you will need 2 confluent 10 cm dishes
of HEK293 cells, 1 dish of unlabeled (light), and one dish of
cells labeled with Arg(13C615N4)/Lys(13C615N2) (heavy). Seed
5 × 106 cells in each 10 cm dish in culture medium supplemented with either light or heavy labeled amino acids containing 10% magnetite solution (see Note 8). Incubate for 24 h
pulse time at 37 °C, 5% CO2.
5.After pulse time, aspirate the medium containing magnetite
and carefully wash the cells thrice with PBS (see Note 9).
6.After washing, add regular culture medium for a 24–36 h
chase.
7. Treat your cells with the desired stimulus prior to isolation (see
Note 10).
8. Before starting the isolation procedure, cool your cells, as well
as the isolation buffer and PBS, on ice.
9.Wash cells twice with PBS to remove proteins found in the
culture medium.
10.Add 2 ml of isolation buffer to each 10 cm dish and detach
cells using a cell scraper. Light and heavy labeled cells should
have the same cell density and can be merged at this point. If
cell densities are evidently different, refer to Note 11.
11.Homogenize the cell suspension in a tight-fitting Dounce
homogenizer for 25 strokes placing it in ice water (see Note 5).
12. Transfer homogenate to a 15 ml Falcon tube. Pellet nuclei and
unbroken cells at 600 × g for 10 min at 4 °C.

13. Transfer the postnuclear supernatant (PNS) to a fresh tube and
keep it on ice. Resuspend the cell/nuclear pellet in 4 ml isolation buffer, transfer back into the Dounce homogenizer and
repeat the homogenization procedure. After centrifugation,
merge both supernatants.
14. Insert the LS column into the magnetic stand and add 1 ml of
BSA solution to the column. Let the column empty by gravity
flow.
15.Apply combined postnuclear supernatants (input) to the column and let it pass by gravity flow. Collect non-bound material
(flow-through).
16.Add 1 ml DNAse solution to the column and incubate for
10 min at 25 °C.
17. Wash the column with 5 ml of isolation buffer.
18.Remove the column from the magnetic stand. It may be that
the iron beads of the column retain a certain amount of magnetic field prohibiting efficient elution of the lysosomes. In
order to remove this residual magnetism hit the column against
a hard surface before elution.


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Melanie Thelen et al.

19.Retrieve the lysosomal fraction by adding 500 μl of isolation
buffer to the column and eluting using the plunger. Repeat
this procedure three times (see Note 12).
20.Remove and save around 250 μl of each fraction (input,
flowthrough, wash, and eluate) to determine the
β-hexosaminidase activity in all fractions after the isolation of
lysosomes has been completed.
3.2 

β-Hexosaminidase
Enzyme Assay

To control for efficacy and quality of lysosome enrichment, the
activity measurement of β-hexosaminidase is routinely performed
after each experiment. Its activity serves as indicator for the quantity and integrity of lysosomes during the enrichment procedure. A
representative result for the isolation of lysosomes from HEK293
cells is displayed in Fig. 2. To determine how much of the enzymatic activity is contained in intact lysosomes, the assay is performed for each sample with and without detergent. The difference
between both values equals the activity that is present in intact
lysosomes and is therefore available for purification.
1. Pipet 4 × 25 μl of each fraction into a microtiter plate.
2. Add 2 μl of 10% Triton X-100 to two of the samples.
3. Add 50 μl of substrate solution.
4.As blank, mix 25 μl of isolation buffer with 50 μl of substrate
solution.
5.Incubate for 15 min at 37 °C. Depending on the amount of
cells used incubation time can be prolonged (up to 24 h).
6.Stop the reaction by addition of 200 μl of stop solution. The
β-hexosaminidase-containing samples should turn yellow during this step.
7. Measure absorbance at 405 nm.
8. Calculate enzymatic activity using the Lambert–Beer law.
9. When your isolation is not successful, please see Notes 6 and 13.

Fig. 2 Efficiency of lysosomal enrichment. (a) The total activity of β-hexosaminidase was determined in each
fraction by an enzymatic activity assay with (white bar) and without (grey bar) Triton X-100. Lines next to the
columns represent the portion of intact lysosomes. (b) Western blot image from 0.25% of each fraction.
Membranes were probed with antibodies against cathepsin D (lysosomal lumen), LAMP2 (lysosomal membrane), protein disulfide isomerase (endoplasmic reticulum), Tom20 and VDAC1 (mitochondria)


Isolation and MS-Analysis of Lysosomes


3.3  Mass
Spectrometry Sample
Preparation

9

To avoid contamination of the sample with keratin or other contaminants, always use new plastic ware, pre-stacked tips and MS
grade solvents and chemicals. During work, gloves should be
always worn and frequently exchanged. To avoid the loss of sample, use maximum recovery tubes and pipet tips.
1.The protein concentration of the lysosomal fraction is frequently low (around 0.15–0.4 mg/ml). To concentrate the
eluate, centrifuge in a centrifugal filter unit until you reach
around 10% of the original volume (usually around 100 μl).
2.Add Laemmli buffer to 1× concentration and denature the
samples at 95 °C for 5 min (see Note 14).
3. Cool samples to 25 °C and add 40% acrylamide to a final concentration of 1% (w/v). Incubate at 25 °C for 30 min for alkylation of cysteines (see Note 15).
4. Separate your sample by SDS-PAGE.
5. Wash the gel thrice for 5 min with deionized water on a shaker.
6.Stain your gel with PageBlue™ solution for several hours or
overnight and destain with deionized water.
7. Cut the gel lane in ten equal slices and cut each gel slice subsequently in ~1 mm3 pieces with a scalpel (see Note 16). Transfer
pieces into separate microcentrifuge tubes, one for each slice.
If you do not immediately proceed with the in-gel digestion,
cover the gel pieces with water to avoid drying.
8.Add 500 μl solution A to each fraction, if the gel pieces were
stored in water remove it first.
9. Incubate for 30 min at 25 °C and 1000 rpm in a thermomixer
(see Note 17). Remove and discard the liquid and repeat step
9 twice or thrice until all gel pieces are colorless (see Note 18).
10. Add 500 μl 100% ACN and incubate for 15 min at 25 °C and

1000 rpm in a thermomixer. The gel pieces should turn white
during this step.
11.Remove and discard the liquid and dry the gel pieces in a vacuum centrifuge, this takes typically between 5 and 15 min.
12. Add 10 μl trypsin solution to each sample and wait until the
liquid is soaked up by the dry gel pieces.
13. Add 50 μl of 100 mM NH4HCO3, pH 7.8 and incubate for
10 min at 25 °C.
14. If the gel pieces are not completely covered by liquid, add 100
mM NH4HCO3, pH 7.8 until they are covered.
15. Incubate the samples overnight at 37 °C (see Note 19).
16.Transfer the supernatant of each sample, which contains your
digested peptides, to a fresh microcentrifuge tube (see Notes
20 and 21).


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Melanie Thelen et al.

17. Add 50 μl of solution B to the gel pieces and incubate for
15 min at 25 °C and 1000 rpm in a thermomixer. Transfer the
supernatants to the microcentrifuge tubes from step 16.
18. Add 50 μl of 100 mM NH4HCO3  pH 7.8 and incubate for
15 min at 25 °C and 1000 rpm in a thermomixer. Do not
remove the liquid at the end of this step.
19. Add 100 μl 100% ACN to the samples and incubate for 15 min
at 25 °C and 1000 rpm in a thermomixer. Transfer the supernatants to the microcentrifuge tubes from step 16.
20. Dry the samples using a vacuum centrifuge.
21. Resuspend your samples in 20 μl of solution C.
22. Add 20 μl of Methanol (MeOH) to your Stage Tips and centrifuge at 5000 × g for 30 s. If the liquid does not pass completely, increase centrifugation time accordingly.

23. Add 20 μl of 0.5% (v/v) acetic acid 80% ACN and centrifuge
at 5000 × g for 30 s.
24. Add 20 μl 0.5% (v/v) acetic acid and centrifuge at 5000 × g for 30 s.
25. Apply each sample to one StageTip and centrifuge at 5000 × g
for 30 s.
26.Wash Stage tips with 20 μl 0.5% (v/v) acetic acid and centrifuge at 5000 × g for 30 s.
27. Transfer Stage tips to fresh microcentrifuge tubes (see Note 22).
28. Elute sample with 2 × 20 μl of 0.5% (v/v) acetic acid 80% ACN
by centrifugation for 30 s at 5000 × g and dry the eluate using
a vacuum centrifuge.
29.Resuspend samples in 20 μl of solution C, sonicate in an ultrasonic water bath for 5 min and centrifuge at 20,000 × g for
15 min. Load 5 μl of this solution to an autosampler vial and
proceed to MS analysis. Take the sample from the top in order to
avoid small particles which may have accumulated at the bottom.
3.4  LC-MS-MS
Measurement

1.Load 5 μl of sample on the analytical column using a nanoLC
system (e.g. Thermo EASY-nLC 1000).
2.Load with 100% A at a flow rate of 1 μl/min followed by a
washing step for 10 min with 100% A at a flow rate of 1 μl/min.
3.Elute with a linear gradient from 100% A to 65% A/35% B in
60 min.
4. Set positive ion mode and a capillary voltage of 1800 V for the
ionization of peptides eluting from the column.
5.Acquire survey scan at a mass range m/z 400 to m/z 1200
and a resolution of 60,000 in the Orbitrap mass analyzer, followed by fragmentation of the ten most abundant ions in the
ion trap part of the instrument.
6. Set the repeat count to one and the dynamic exclusion window
to 60 s.



Isolation and MS-Analysis of Lysosomes

3.5  Data Analysis

11

The analysis of the raw files is performed with Proteome Discoverer
(Thermo Scientific) using the MASCOT search engine (www.
matrixscience.com) using databases from www.uniprot.org.
Alternatively, other programs like MaxQuant [30] can be used.
1. Set propionamide at cysteine residues as fixed modification.
2.Set as variable modifications: protein N-acetylation, methionine oxidation, isotopic labeling of arginine (13C615N4) and
lysine (13C615N2) and N-terminal conversion of glutamic acid
and glutamine to pyroglutamic acid.
3. Set accepted missed cleavages to two. Here a mass tolerance of
10 ppm for the precursor ion and 0.6 Da for the fragment ions
were applied, but these properties should be adjusted to the
performance of the mass spectrometer used.
4.As a quantification method, use SILAC 2plex and normalize
results to mean protein amount.
5. Process search results with a false discovery rate of 0.01 and only
consider proteins with at least two unique peptides and peptide
spectral matches with high confidence for quantification.
We usually recover ~80% of intact lysosomes in the lysosomal
eluate fraction as determined by β-hexosaminidase assay (Fig. 2a). In
the corresponding mass spectrometric analysis, however, we identify
about 3000 proteins. Western blots for marker proteins covering
lysosome, endoplasmic reticulum, and mitochondria show that the

eluate of the magnetic column supposedly contains solely lysosomes
and is free of non-lysosomal proteins (see Fig. 2b) as the respective
marker proteins, when comparing equal volume percentages of each
fraction, are only detected in the input but not in the eluate fraction.
This is apparently not the case, otherwise we would only detect the
lysosomal proteins (see Table 1) and nothing else. One must keep in
mind, however, that the high sensitivity of modern mass spectrometers can exceed that of a Western blot and that the samples used for
mass spectrometric analysis are concentrated using spin filters.
Therefore, it is not unusual that the majority of proteins identified in
a mass spectrometric dataset generated from such samples are contaminating non-lysosomal proteins from all cellular compartments.
Therefore, in order to evaluate the suitability of a lysosomal
enrichment procedure for mass spectrometric analysis, one should
not only consider the number of non-lysosomal proteins but rather
how many lysosomal proteins can be detected in the fraction of
enriched lysosomes. To assess this for the method described here,
we prepared a list containing proteins which are currently verified
to be of lysosomal localization (see Table 1). Some of these proteins
may be specific to a cell type or a cellular condition. In a routine
experiment we detect 136 of 186 verified lysosomal or lysosome-­
associated proteins corresponding to a coverage rate of ~73%. The
high number of non-lysosomal proteins in our routine data sets
makes it difficult to determine whether proteins which have so far


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Melanie Thelen et al.

Table 1
Overview of all proteins with experimentally verified exclusive or partial lysosomal localization

Uniprot ID

Gene

Q9BZC7

ABCA2

Q9NP58

ABCB6

Q9NP78

ABCB9

O14678

ABCD4

P11117

ACP2

P13686

ACP5

P20933


AGA

Q96B36

Identified

Uniprot ID

Gene

Q9UJQ1

LAMP5

Q6IAA8

LAMTOR1

×

Q9Y2Q5

LAMTOR2

×

×

Q9UHA4


LAMTOR3

×

×

Q0VGL1

LAMTOR4

×

O43504

LAMTOR5

×

Q15012

LAPTM4A

×

AKT1S1

Q86VI4

LAPTM4B


P28039

AOAH

Q13571

LAPTM5

P02743

APCS

P17931

LGALS3

Q96BM9

ARL8A

×

O00214

LGALS8

Q9NVJ2

ARL8B


×

Q99538

LGMN

P15289

ARSA

×

Q99732

LITAF

P15848

ARSB

×

P38571

LIPA

Q96EG1

ARSG


Q9NUN5

LMBRD1

Q6UWY0

ARSK

×

Q969J3

LOH12CR1

×

Q13510

ASAH1

×

O00754

MAN2B1

×

Q15904


ATP6AP1

×

Q9Y2E5

MAN2B2

×

Q93050

ATP6V0A1

×

O00462

MANBA

×

P27449

ATP6V0C

×

Q5T0T0


MARCH8

P61421

ATP6V0D1

×

Q9GZU1

MCOLN1

O15342

ATP6V0E1

Q96EZ8

MCRS1

P38606

ATP6V1A

×

Q96FH0

MEF2BNB


×

P21281

ATP6V1B2

×

Q9H3U5

MFSD1

×

P21283

ATP6V1C1

×

Q8NHS3

MFSD8

×

Q9Y5K8

ATP6V1D


×

Q9BVC4

MLST8

×

P36543

ATP6V1E1

×

P05164

MPO

Q16864

ATP6V1F

×

P42345

MTOR

×


O75348

ATP6V1G1

×

Q02083

NAAA

×

Q9UI12

ATP6V1H

×

P17050

NAGA

×

Q6UW56

ATRAID

×


P54802

NAGLU

×

Q07812

BAX

×

Q92542

NCSTN

×

×

×

Identified

×

×

×


(continued)