PhD thesis Accessory Proteins at ERES-

FACULTY OF SCIENCE
UNIVERSITY OF COPENHAGEN
PhD thesis
David Klinkenberg
Accessory Proteins at ERES‐
Assembly of ER exit sites is regulated by interactions of p125A with lipid signals.
Academic advisor: Lektor Lars Ellgaard, PhD
Co‐Supervisor: Assoc. Prof. Meir Aridor, PhD (University of Pittsburgh)
This thesis has been submitted to the PhD School of The Faculty of Science, University
of Copenhagen: 26/02/2013

Name of department: Department of Biology
Author: David Klinkenberg
Title / Subtitle: Accessory Proteins at ERES‐
p125A Couples Lipid Signals with Functional ER Exit Site Assembly
Subject description: This thesis provides a characterization of the accessory protein p125A and its
functions at Endoplasmic Reticulum Exit Sites (ERES) in response to membrane
lipid composition by dissecting two functional domains within p125A. The
results provide evidence for a mechanism where p125A response to lipid
signals, i.e. PI(4)P, promotes both COPII displacement from the scaffolding
protein mSec16A, as well as stabilizing linkage between the two layers
forming the COPII cage at ERES.
Academic advisor: Lektor Lars Ellgaard, PhD
Co‐Supervisor: Assoc. Prof. Meir Aridor, PhD (University of Pittsburgh)
Submitted: 26 February 2013
Grade: PhD
Front Page Image
EGFPp125A expression co-localized at ERES
(yellow) with Sec31A (green) at 10°C.
Recorded on a Olympus Fluoview 1000 PLAPON
60 x objective, NA = 1.42

Abstract
Traffic mediated by vesicles budding from the membranes of the Endoplasmic Reticulum (ER) at
specific sites termed ER Exit Sites (ERES) is mediated by the COPII machinery. The molecular
1
interactions that COPII utilize to form the basic bud have been examined and mapped. However, not
much is known about how these interactions are regulated, in particular with respect to COPII
interactions in relation to specific ER membrane lipid signals.
This thesis explores the mechanisms by which the accessory protein p125A (aka Sec23IP) regulates
COPII at ERES. The work shows that p125A recognizes lipids, and in particular phosphatidylinositol‐4‐
phosphate (PI(4)P), through concerted actions between two internal domains – a sterile α‐motif
(SAM) and a putative lipid recognizing domain termed a DDHD domain. We demonstrate that p125A
binding at ERES, in response to local presence of PI(4)P, mediates displacement of the two COPII
layers from the Sec16A ERES nucleation scaffold. We furthermore show evidence that p125A
provides a linkage between the two COPII layers during vesicle budding. Additional observations
indicate that p125A lipid recognition and binding supports the steady‐state transport levels between
ER and Golgi. We also provide evidence that Sec16A functions at an early stage of ERES assembly, as
we can show clear segregation of Sec16A from ERES during temperature imposed inhibition of the
cellular transport.
We finally explore the membrane binding mechanism of mammalian Sec16 (mSec16) A and B, and
identify domains within each mSec16 subtype that show membrane binding, but do not support
selective ERES targeting.
A major part of the experimental work presented in this thesis is included in the following
manuscript that has been submitted for review to the Journal of Cell Biology:
Assembly of ER exit sites is regulated by interactions of p125A with lipid signals.
David Klinkenberg, Kimberly R. Long, Kuntala Shome, Simon C. Watkins and Meir Aridor
26/2‐2013
David Klinkenberg

Acknowledgments
The experiments of this thesis were all performed in the laboratory of Ph.D. Meir Aridor at the
Department of Cell Biology, University of Pittsburgh, Pittsburgh PA, USA, while employed as
Research Technician.
I would first like to thank Meir for all his encouragement and long scientific discussions that have
2
fueled my fascination for this particular field of Cell Biology, and in particular for allowing me to turn
a position as Research Technician into a Ph.D. project.
I would also like to give a very special thanks to Ph.D. Kimberley Long for helping verify and finalize
the results of the appended manuscript, Ph.D. Kuntala Shome for providing technical assistance, and
the rest of the members of the Aridor lab, past and present, that I had the great pleasure to work
with while in Pittsburgh and whom have made me grow as a scientist.
I would finally like to thank my mother Marta and my step‐dad Carsten for all their support and aid
that made it possible for me to move for an extended period to the beautiful city of Pittsburgh,
thereby giving me the opportunity to conduct the research presented in this thesis.
In memoriam Teresa.

Summary
The components of the COPII machinery, which are essential in establishing an effective
Endoplasmic Reticulum (ER) to Golgi transport from ER exit sites (ERES), have been identified and
characterized within the last 25 years. These consist of the essential Sec12, Sec23, Sec24, Sec13,
Sec31 and Sar1 proteins. Together these components co‐operate in cargo‐selection as well as
forming, loading and releasing budding vesicles from specific regions on the membrane surface of
the ER. Coat components furthermore convey vesicle targeting towards the Golgi. However, not
much is known about the mechanisms that regulate the COPII assembly at the vesicle bud site.
This thesis provides the first regulatory mechanism of COPII assembly in relation to ER‐membrane
lipid‐signal recognition by the accessory protein p125A (Sec23IP).
The aim of the project was to characterize p125A function by dissecting two main domains in the
protein; a putative lipid‐associating domain termed the DDHD domain that is defined by the four
3
amino acid motif that gives the domain its name; and a ubiquitously found domain termed Sterile α‐
motif (SAM), which is mostly associated with oligomerization and polymerization.
We first show, that the DDHD domain of p125A utilizes a stretch of positively charged residues
(KGRKR) to bind lipid membranes that are enriched in Phosphatidylinositol‐4‐phosphates (PI(4)P).
The specificity of the DDHD domain lipid recognition is demonstrated to be enhanced through p125A
oligomerization mediated by the upstream SAM domain.
We then show that p125A is targeted specifically to ER exit sites (ERES) through a series of
experiments where p125A expressing cells are incubated at lower temperatures. Incubation at either
15°C or 10°C inhibits cargo transport out of specific compartments that represent defined stages
during the biosynthetic transport between the ER and the Golgi. We find that p125A associates
predominantly with COPII‐marked ERES and dissociates from both the ER‐to Golgi‐intermediate‐
compartment (ERGIC) and from the cis‐Golgi compartment.
The same set of experiments also provides evidence that p125A functions at a later stage of the ER
export. The temperature‐dependent block of ER export is shown to cause a clear segregation of ERES
composed of Sec31A, Sec23 and p125A from the known COPII‐associating ERES nucleation scaffold
protein mSec16A. The temperature block furthermore causes mSec16A to collect on the ER
membrane in structures that neither co‐localize with ERGIC nor Golgi.

Using p125A double mutants that are impaired in lipid recognition, we show that the lipid
recognizing activity of p125A regulates COPII organization. These double mutants are produced by
4
introducing a point mutation (L690E) in the SAM domain that causes inhibition of its oligomerization,
combined with either a charge reversal of the KGRKR lipid recognition motif within the DDHD
domain (850(KGRKR/EGEEE)854 – DDHD‐PI‐X) or by deleting the entire DDHD domain (ΔDDHD). We
demonstrate that p125A double mutants with defective lipid recognition strongly disperse ERES. This
dispersal of the ERES can be rescued by replacing the DDHD with the PI(4)P recognizing Fapp1‐PH
domain even if SAM(L690E) is still present in p125A. We additionally show that a stretch of cationic
residues (KGRKR) in the DDHD abrogated p125A lipid recognition influences the proteins residency
time at ERES.
Comparison of overexpressed of p125A wt, p125A(L690E)(PI‐X) and p125A(L690E)(ΔDDHD) with the
expression of a GFP‐tagged mSec16A provides evidence that p125A lipid recognition furthermore
promotes the displacement of COPII from the mSec16A scaffold during ERES assembly. The
overexpression of p125A wt and p125A(L690E)(ΔDDHD), but not p125A(L690E)(PI‐X), causes p125A
to aggregate in enlarged structures. The enlarged p125A wt structures show clear segregation from
mSec16A, whereas the enlarged p125A(L690E)(ΔDDHD) structures become engulfed by the
mSec16A. Surprisingly, no inhibition in the overall export of the temperature sensitive VSV‐G
transport marker can be measured during these conditions.
Depletion of p125A by RNAi is additionally shown to cause perturbation of steady state level
transport in HeLa cells. The transport perturbation manifests itself by the dispersion/shattering of
the Golgi ribbon, where the Golgi instead appears to be broken into multiple mini‐stacks adjacent to
ERES. The steady state transport level can be rescued by the introduction of an RNAi resistant p125A
wt clone, but not by an RNAi resistant p125A double mutant.
These findings taken together point towards a model of p125A regulation at ERES, where p125A
association with Sec31A, Sec23 and to specific ER membrane lipid signals provides linkage between
the two COPII layers, and furthermore promotes displacement of the COPII cage from the mSec16A
scaffold.
We additionally identify a structural fold termed WWE in the unstructured region of the p125A N‐
terminus that may potentially promote p125A binding to Sec31A.
We then further expand the temperature dependent ER export analysis of mSec16A to its smaller
homolog mSec16B. Here, we examine mSec16B and mSec16A with regards to both proteins
membrane targeting and association with ERES. We determine the localization of Sec16B by

5
transient expression in HeLa cells, and find that the protein is evenly distributed throughout the cell
except the nucleus at 37°C, as is also observed with mSec16A. When the temperature is lowered to
15°C, mSec16B mimics mSec16A further by associating and forming larger defined structures at the
ER membrane that do not co‐localize with COPII, ERGIC53 or cis‐Golgi. Lowering the temperature
further to 10°C, which arrests cargo at the ERES, maintains the formed structures substantially and
decreases the even cellular distribution of mSec16B.
We further dissect both mSec16A and mSec16B, and show that the region in human mSec16B
encompassing residues 35‐194 and the region in human mSec16A comprising residues 1096‐1190
maintain membrane binding irrespective of the removal of membrane associating proteins by salt
wash or proteolytic digestion. However, neither mSec16B (35‐194) nor mSec16A (1096‐1190)
maintain ERES targeting.
These findings support previous observations of the need for the membrane binding regions to be
expressed in cis with a Central Conserved Domain (CCD) in both proteins to convey ERES targeting.

Dansk Resumé (Summary in Danish)
6
De komponenter, der er essentielle for etableringen af en effektiv Endoplasmatisk Reticulum (ER)‐til‐
Golgi transport, er blevet identificeret og karakteriseret indenfor de sidste 25 år. De udgøres af
proteinerne Sec12, Sec23, Sec24, Sec13, Sec31 og Sar1, der samarbejder ved sorteringen af cargo,
samt former, laster og afsnører vesikler fra særlige regioner på membranoverfladen af ER, hvor de
endvidere sørger for, at vesiklerne målrettes henimod Golgi. Desværre ved man meget lidt om de
mekanismer, som regulerer COPII ved ”bud sitet” for vesikler.
I denne afhandling giver vi for første gang en beskrivelse af en reguleringsmekanisme for COPII
samling, der er varetaget af "accessory" proteinet p125A (Sec23IP) ved hjælp af dets evne til at
genkende særlige lipid‐signaler i ER‐membranen.
Formålet med dette projekt har været at karakterisere p125A’s funktion ved at dissekere to
hoveddomæner i proteinet: Et formodet lipidbindende domæne, der defineres af et 4‐aminosyre‐
motiv (DDHD domænet), samt et oligomeriserings‐domæne kaldet Sterile α‐Motif (SAM), som findes
i en række multidomæneproteiner, og som endvidere oftest er tilknyttet oligomerisering og
polymerisering.
Vi viser først, at p125As DDHD‐domæne igennem et positivt ladet motiv (KGRKR) interagerer med
lipidmembraner, der er beriget med phosphatidylinositol‐4‐phosphater (PI(4)P). Specificiteten for
DDHD‐domænets lipidgenkendelse forstærkes gennem p125A's oligomeriseringen medieret af det
opstrøms SAM‐domæne.
Dernæst viser vi, at p125A hovedsagligt forefindes ved ER exit sites (ERES). Ved at inkubere celler,
der udtrykker p125A, ved forskellige temperaturer lavere end 37C, hæmmes cargo‐transporten ud
af specifikke compartments. Disse compartments repræsenterer hver især forskellige stadier af den
biosyntetiske transport. Disse eksperimenter viser, at p125A især lokaliserer til ERES. Desuden viser
vi, at p125A hovedsagligt associerer med COPII‐markerede ERES og dissocierer fra både "ER‐to‐Golgi‐
intermediary compartments" (ERGIC) og fra cis‐Golgi.
Den samme eksperimentrække antyder også, at p125A fungerer under et senere stadium af ER
eksporten. Den temperaturafhængige blokering af ER eksport medfører en klar adskillelse af ERES
bestående af Sec31A, Sec23 og p125A fra mSec16A, der er et kendt ERES dannende "scaffold"

protein. Endvidere medfører den temperaturafhængige blokering til, at mSec16A samles på ER
membranen i strukturer, der ikke co‐lokaliserer med hverken ERGIC eller Golgi.
Gennem brugen af p125A dobbeltmutanter, der er hæmmede i deres evne til at genkende lipider,
påviser vi, at p125A's lipidgenkendelse er med til at regulere COPII organisationen. De pågældende
dobbeltmutanter er skabt ved at introducere en punktmutation (L690E) i SAM domænet, der
inhiberer domænets evne til oligomerisere, kombineret med enten en positiv til negativ
7
ladningsændring i en strækning af aminosyrer i DDHD domænet (850(KGRKR/EGEEE)854 – DDHD‐PI‐
X), eller ved helt at fjerne DDHD domænet igennem en deletion (ΔDDHD). ERES spredes som
konsekvens af den introducerede hæmning af p125A's lipidgenkendelse. Spredningen af ERES kan
reddes ved at udskifte DDHD domænet i p125A med det PI(4)P genkendende Fapp1‐PH domæne,
også under indflydelse af SAM(L690E). Vi påviser ydermere, at den hæmmede lipidgenkendelse har
indflydelse på p125A's opholdstid ved ERES.
Sammenligning af overudtrykt p125A wt, p125A(L690E)(PI‐X) og p125A(L690E)(ΔDDHD) i forhold til
GFP‐mærket mSec16A antyder, at p125A's lipidgenkendelse også fremmer COPII's afkobling fra
mSec16A's "scaffolding" ved ERES dannelsen. Overudtrykket af p125A wt og p125A(L690E)(ΔDDHD),
men ikke p125A(L690E)(PI‐X), fører til, at p125A aggregerer i større strukturer. Der ses en tydelig
adskillelse imellem de forstørrede p125A wt strukturer og mSec16A, hvorimod de forstørrede
p125A(L690E)(DDHD) strukturer til gengæld lader til at være fuldstændigt opslugt af mSec16A. Til
vores overraskelse lader den tilstedeværende ER eksport til ikke at være hæmmet nævneværdigt,
når den måles ved hjælp af transportmarkøren VSV‐G.
Vi viser også, at nedregulering af p125A ved RNAi forårsager en kraftig forstyrrelse af steady‐state
niveauet for transporten i HeLa celler, hvilket manifesterer sig i spredning ("shattering") af Golgi
"ribbon", der i stedet bliver nedbrudt til små mini‐stacks overfor ERES. Steady‐state transporten kan
reddes ved introduktionen af et RNAi‐modstandsdygtigt p125A wt‐konstrukt, men ikke af en RNAi‐
modstandsdygtig dobbeltmutant ‐ p125A (L690E)(PI‐X).
Samlet peger disse observationer på en model af p125A's regulering ved ERES, hvor p125A
associering med Sec31A, Sec23 og til særlige lipidsignaler i ER‐membranen yder en form for kobling
imellem det indre og det ydre lag af COPII, og samtidig også sørger for at COPII "cagen" afkobles fra
mSec16A "scaffoldingen".
Derudover, identificerer vi et strukturelt fold kaldet et WWE domæne, der befinder sig i en N‐
terminal ustruktureret region af p125A, og som har potentiale for at formidle p125A’s binding til
Sec31A.

Vi udvider endvidere analyser af den temperatur‐afhængige blokering af ER eksporten til også at
omhandle mSec16A's mindre homolog mSec16B. Vi undersøger først lokaliseringen af Sec16B ved
transient udryk i HeLa celler og finder, at ved 37°C udtrykkes proteinet spredt udover det meste af
8
cellen bortset fra cellekernen, hvilket også er tilfældet med mSec16A. Sænkes temperaturen til 15°C,
arter mSec16B sig videre som Sec16A og associerer kraftigt med membraner, hvor mSec16B samler
sig til større definerbare strukturer ved især ER‐membranen. De observerede strukturer co‐
lokaliserer ikke med COPII, ERGIC53 eller cis‐Golgi. Yderligere sænkning af temperaturen til 10°C,
hvilket forårsager en blokering for transport af cargo ud af ERES, bibeholdes de pågældende
strukturer med en drastisk reduktion i mængden af mSec16B, der før var jævnt fordelt ud over
cellen.
Dernæst dissekerer vi både Sec16A og Sec16B i mere detalje. Vi påviser herved at regionen i Sec16B
omfattende aminosyrerne 35‐194, samt regionen i Sec16A omfattende aminosyrerne 1096‐1190,
bibeholder membranbinding uanset om man fjerner membranassocierede proteiner ved enten
saltvask eller proteolytisk fordøjelse. Derimod bibeholder hverken Sec16B (35‐194) eller Sec16A
(1096‐1190) målretningen imod ERES.
Disse resultater bekræfter forudgående observationer med hensyn til behovet af, at de
membranbindende regioner i Sec16A og Sec16B skal udtrykkes i cis med et centralt konserveret
domæne (CCD) i begge proteiner for at formidle målretning til ERES.

Table of Contents
Abstract 1
Acknowledgements 2
Summary 3
Dansk Resumé (Summary in Danish) 6
Table of Contents 9
9
Abbreviations 13
Introduction: 16
The Discovery of two Organelles and the Link Between Them 16
The Biosynthetic transport pathway: a brief overview 17
The Endoplasmic Reticulum (ER) and the ER‐to‐Golgi intermediate compartment – ERGIC 20
ER dynamics, morphology and general function 20
ER exit sites and the ERGIC 21
The ERGIC53 protein 23
The Golgi apparatus and COPI 23
General mammalian Golgi morphology 24
Golgi and cisternal maturation 25
COPI 25
COPI cargo loading 27
COPII 27
Sec12 28
Sar1 29
Sec23 and Sec24 31
Sec13 and Sec31 32

10
Cargo loading end ER export motifs 35
COPII mutations and physiological effects 36
Membranes and lipid biogenesis 38
Lipid transport 39
Cholesterol and membrane fluidity 40
Lipids and membrane curvature 41
PI and phosphorylated PI (PIP): their role in signaling 43
Golgi and PI(4)P 45
PI(4)P and ERES formation 45
Sec16 48
Sec16 structure 49
Sec16 functions 50
Sec16B 53
p125A (Sec23IP) 54
p125A architecture 54
p125B 55
Cellular localization of p125A 57
Consequences of modulating p125A expression levels 57
P125A ERES targeting and interactions 58
p125A and disease 60
References 61
Aim of the Project 80
Publication with Supl.: Assembly of ER exit sites is regulated by interactions of p125A with lipid signals 81
Abstract 82
Introduction 83

11
Results 85
p125 is recruited with COPII to PI4P enriched liposomes 85
The DDHD and SAM domains cooperate to support lipid recognition in vitro and
binding of PI4P‐rich membranes in cells 86
Segregation of ERES from ERGIC and Golgi at low temperatures reveals and exclusive
localization of p125A at ERES 89
COPII‐p125A containing ERES segregate from mSec16A at low temperatures 90
Charge and hydrophobic interactions are used by the SAM and DDHD domains to
support lipid recognition and assembly 91
Assembly controlled lipid‐recognition is required to regulate COPII organization at ERES 92
Lipid recognition controls p125A residency at ERES 94
p125A functions at a late stage in ERES nucleation 95
Functional contribution of the SAM‐DDHD membrane‐binding module 97
Discussion 98
The SAM‐DDHD lipid‐binding module 98
Role of p125A in ERES regulation 100
Materials and Methods 103
Acknowledgment 109
Abbreviations 109
References 110
Figure legends 114
Supplement (Legends) 120
Figures (with Supplemental figures) 122
Investigations of p125A‐Sec31A associations and mammalian Sec16A and B membrane binding 135
Additional exploration of p125A 135
A study of Sec16A and B membrane binding 141

12
Materials and Methods 152
References 155
Conclusions, Discussion and Perspectives 157
Summary of findings 157
SAM – a domain for oligomerization 159
DDHD Domains and the influence of lipid recognition 161
WWE domain of p125A – a possible Sec31A binding motif 163
Sec16A and B collect into structures at low temperature incubation 167
p125A mediated displacement of Sec16A from ERES 169
Sec16A and Sec16B membrane binding and ERES targeting 173
Physiological Relevance of p125A Regulation 174
Concluding remarks 176
References 178
Co‐authorship Statement 182

Abbreviations
13
ACE 1 – Ancestral Coatomer Element 1
ADP – Adenosine di‐phosphate
Alg‐2 – Alix linked gene ‐2
AP – adaptor protein
Arf – ADP ribosylation factor
ArfGAP – Arf GTPase Activating Protein
ATP – Adenosine tris‐phosphate
BFA – Brefeldin A
CCD – Conserved Central Domain
CDP – Cytosine di‐phosphate
Cer – Ceramide
CERT ‐ Ceramide Transfer protein
CMP – Cytosine mono‐phosphate
co‐IP – co‐immuno‐precipitation
COP – Coat Protein
DAG – diacylglycerol
DAGKδ – diacylglycerol kinase δ
DNA – deoxy‐ribonucleic acid
DRM – detergent resistant membrane
DsRNAi – Dicer specific ribonucleic acid inhibition
ECFP – enhanced cyan fluorescent protein
EGFP – enhanced green fluorescent protein
EH – end‐helix
EM – electron microscopy
ER – Endoplasmic Reticulum
ERAD – ER associated degradation machinery
ERES – ER Exit Sites
ERGIC – ER‐Golgi intermediary compartment
ERK7 – Extracellularly Regulated Kinase 7
ER‐RLM – endoplasmic reticulum derived rat liver microsomes
ERv – ER‐Vesicle protein
EYFP – enhanced yellow fluorescent protein
FAPP – Four‐Phosphate‐Adaptor Protein
FRAP – fluorescence recovery after photobleaching
G3P – glycerol‐3‐phosphate
GalNAc – N‐acetylgalactosaminyltransferase
GAP – Guanosine activating protein
GAT1 – GABA transporter 1
GDP – Guanosine di‐phosphate
GEF – Guanosine Exchange Factor
GFP – green fluorescent protein

14
GGA's – Golgi‐Localized γ‐ear containing, Arf‐binding proteins
GPI – glycosylphosphatidylinositol
GST – glutathione transferase
GTP – Guanosine tris‐phosphate
HeLa – Henrietta Lacks
kDa – kilo Dalton
KDELR – KDEL recognizing receptor
LPA – lysophosphatidic acid
LPAT – lysophosphatidic acid transferase
mAB – monoclonal antibody
ML – mid‐loop
mRFP – monomeric red fluorescent protein
mRNA – messenger ribonucleic acid
MT – microtubule
MTOC – microtubule organizing center
MVB – Multivesicular Body
MW – molecular weight
Nir – N‐Terminal domain interacting receptor
NM – Nodular Melanoma
NRK – newborn rat kidney
PA – phosphatidic acid
PA‐PLA1 – phosphatidic acid preferring‐Phospholipase A1
PAR – poly(ADP)‐ribosylation
PARP – poly(ADP)‐ribosylation protein
PC – phosphatidylcholine
PCR – polymerase chain reaction
PE – phosphatidylethanolamine
Pex – Peroxisome specific transport receptor
PG – phosphatidylglycerol
PH – pleckstrin homology
PI – phosphatidylinositol
PI(3)P – phosphatidylinositol‐3‐phosphate
PI(3,5)P2 – phosphatidylinositol‐3,5‐bis‐phosphate
PI(4)P – phosphatidylinositol‐4‐phosphate
PI(4,5)P2 – phosphatidylinositol‐4,5‐bis‐phosphate
PI4KinIIIα – phosphatidylinositol‐4 Kinase type III α
PI4KinIIIβ – phosphatidylinositol‐4 Kinase type III β
PI4KinIIα – phosphatidylinositol‐4 Kinase type II α
PIK – Phopsphatidylinositol Kinase
PIP – phosphatidylinositol‐phosphates
PITP – PI transfer domain
PM – plasma membrane
PMA – phorbol 12‐myristate 13‐acetate
P‐Q – proline‐glutamine rich
PS – phosphatidylserine
qPCR – quantitative polymerase chain reaction

ER – rough
RLC ‐ rat liver cytosol
RNA – ribonucleic acid
RNAi – ribonucleic acid inhibition
SAM – Sterile α‐Motif
SEC (Sec) – secretory deficient
SFV – Simliki Forest Virus
siRNA – small inhibitory ribonucleic acid
SNARE – soluble NSF (N‐ethylmaleimide‐sensitive factor) attachment protein (SNAP) receptors
SSM – Superficial Spreading Melanoma
STAM – signal‐transducing adaptor molecule
TAC – Tip Attachment Complex
TEL – translocation ETS leukemia
tER – transitional ER
TFG‐1 – Tyrosine Receptor Kinase Fused Gene‐ 1
TGN – trans Golgi network
TRAPP – Transport/Trafficking Protein Particle
VAP‐A & VAP‐B – Vesicle Associated membrane Protein A & B
VSV‐G – Vesicular Stomatitis Virus Glycoprotein
VSV‐G‐tsO45 – temperature sensitive late phase G‐protein from Vesicular Stomatitis Virus
capsid
VTC – Vesicular Tubular Clusters
WB – Western Blot
15

Introduction
16
This introduction will briefly describe the function and the different steps of the secretory
transport pathway. An overview of some of the stations, organelles, and important
components involved in the different stages of transport will also be given in the first part.
The second part provides a comprehensive review of the actual initiation of the transport at
its origin, the Endoplasmic Reticulum (ER), with particular emphasis on two important
proteins necessary for the transport initiation process, namely Sec16 and p125A.
The discovery of two organelles and the link between them
The discovery of the major components involved in the biosynthetic transport pathway
begins in 1898 when the Italian physician Camillo Golgi was able to visualize a "cellular
body" in Purkinje cells by staining it with silver nitrate [1]. He describes this cellular structure
as: "..ora ha struttura reticolare, ora appare in forma di strato continuo omogeneo, ora si
direbbe costituito da fine squammette applicate in continuità l'una dall'altra..". What he saw
was a net‐like/reticular structure surrounding the nucleus of the Purkinje cell in
homogenous continuous stratas. He calls them "apparato interno reticolare". Later this
organelle gets named "the Golgi structure" in honor of its discoverer [1]. The function of this
cellular body does not become clear until 70 years later when James D. Jamieson and
George E. Palade are able to define the Golgi as a regular way station in protein transport
between the ER and vacuoles [2]. Just prior to this, Marian Neutra and C.P. Leblond were
able to recognize that the Golgi played a role in the synthesis of complex carbohydrates and
glycoproteins in secretory mucosal cell of rats [3, 4].
In 1945 Keith R. Porter, Albert Claude and Ernest F. Fullam discover a second major
component of the transport pathway while examining different types of electron
microscopy (EM) staining procedures on tissue cultures derived from chicken embryos. In
samples stained with osmium they cannot help noticing a "lace‐like reticulum" that extends

throughout the cytoplasm [5]. K.R. Porter later names the network the Endoplasmic
Reticulum [6].
13 years later Philip Siekevitz and George E. Palade noticed that they were able to extract
protein precursors, zymogens, from microsomes that they could identify as mainly being
derived from fractions associated to the rough ER (rER) [7‐9]. In 1960, these researchers
17
finally demonstrated that the ER played an important role in the production of proteins that
were bound for export out of the cell. They injected DL‐leucine‐1‐C 14 into guinea pigs 1 h
after feeding. Through pulse‐chase analysis they were then able to show that the majority of
the digestive protease pre‐cursor chymotrypsinogen was found in rER microsome fractions
from pancreas extracted 1‐3 min post‐injection [10]. In 1964, Lucien G. Caro and George E.
Palade made it possible to map the directional transport in the same pancreatic cells from
guinea pigs by injecting them with DL‐leucine‐ 4,5‐H 3 . They then followed the isotopically
labeled secretory proteins from their translation in the ER, across the Golgi ending up in
discernible vacuoles [11].
Today, we have gained an in‐depth understanding of a variety of functions of both the ER
and the Golgi apparatus, including knowledge about central processes such as protein and
lipid synthesis, protein folding and misfolding, co‐ and posttranslational modification, Ca 2+ ‐
storage, membrane transport and much, much more.
The biosynthetic transport pathway: a brief overview.
The main purpose of the biosynthetic transport pathway is to shuttle newly formed proteins
and lipids to various destinations within and outside of the cell (see fig. 1). The process
begins with the synthesis of protein or lipid at the ER. Export of the newly formed
components out of the ER is initiated by assembly and packaging of the components into
transport vesicles at sites dedicated to vesicle budding. Naturally, these sites have been
named ER exit sites (ERES). The budding of the actual transport vesicles is controlled by an
intricate machinery named the COat Protein (COP) II complex. Vesicle formation starts with
the ER membrane resident protein Sec12 recruiting and initiating the small GTPase Sar1.
Sar1 initiation causes the protein to tether and deform the membrane, and then recruits the

additional components of the COPII machinery Sec23, Sec24, Sec13 and Sec31. Together
18
they form a cage structure that acts as a protein scaffold during the vesicle formation. The
actual COPII cage consist of two protein layers built up of two different heteromeric
complexes; an inner layer consisting of the proteins Sec23 and Sec24, and an outer layer
consisting of the proteins Sec13 and Sec31. The COPII complex is also responsible for
maintaining the transport targeting from the ER towards the Golgi. The transport proceeds
through different way stations where additional sorting and processing occurs.
Figure 1 ‐ ER to Golgi transport pathway ‐ The biosynthetic transport pathway between ER and Golgi. COPII cargo
vesicles are formed from the ER at Vesicular Tubular Structures (VTC) by multiple ER Exit sites. Vesicles are budded in
response to Sec12 recruiting and activating Sar1. Activation of Sar1 in turn recruits the inner layer of COPII,
Sec23/Sec24, where Sec24 aids in loading cargo into the forming vesicle. Next, the outer layer of the cage ‐
Sec13/Sec31 ‐ is recruited followed by membrane fission that releases the budding vesicle. Released COPII vesicles
move anterograde and fuse with themselves and with retrograde COPI vesicles into the ER‐to‐Golgi intermediate
compartment (ERGIC), marked by the lectin ERGIC53. Transport continues from ERGIC to the cis‐Golgi where cargo
enters the Golgi stack for further post‐translational processing. Recycling of ER components is maintained by
retrograde COPI vesicles, that bud off the cis‐Golgi and return to the ER via the ERGIC.
The first stop occurs at a dynamically maintained organelle located between the ER and the
Golgi, the ER‐to‐Golgi intermediate compartment (ERGIC). The main function of the ERGIC
has not yet been fully determined, but it is believed to act as a primary sorting station used
for the retrieval of ER resident factors not destined for the Golgi. From the ERGIC, the
transport proceeds towards and into the Golgi. Within the Golgi, the newly formed proteins
are further processed and matured for their final function. The proteins are conveyed
through the Golgi either within one of the cisternae that form the Golgi, or the proteins are
transported between the individual Golgi cisternae by a Golgi‐dedicated vesicle transport

19
system. The lipids on the other hand become parts of the Golgi membranes where further
modification may occur to prime them for specific tasks, either as signaling molecules or for
altering membrane properties such as fluidity, rigidity or bending.
Transport out of the Golgi finally shuttles the processed proteins or modified lipids to their
site of function through transport in distinct populations of vesicles. These are destined
either for exocytosis by fusion with the plasma membrane (PM), or targeted to intra‐cellular
organelles or compartments by an endocytic vesicle transport system. The initiation of the
transport out of the Golgi is mediated by mechanisms very similar to mechanisms employed
by the COPII machinery. A main difference is that transport out of the Golgi utilizes a
different subset of coat components named clathrin. Clathrin is also used on the PM to
initiate and stabilize vesicle transport processes targeting components in contact with or
from the extracellular environment to compartments within the cell. These vesicles are part
of the endocytic vesicle system mentioned above. The endocytic vesicles target a wide
variety of intracellular organelles, such as endosomes, multivesicular bodies (MVB),
lysosomes, and even return components to the Golgi.
The transport direction away from the ER is generally termed anterograde transport. A
transport system also exists that has directionality towards the ER. Transport in this
direction is termed retrograde transport. An important complex involved in the retrograde
transport is the COPI machinery. COPI functions are quite homologous to the functions of
COPII, the main difference being that COPI vesicle formation mainly occurs on the
membranes of the Golgi. The mechanisms involved in COPI‐mediated vesicle formation are
also very similar to the mechanisms used by COPII. The previously mentioned Golgi‐
dedicated vesicle transport system is believed to mainly consist of COPI coated vesicles.
These are furthermore responsible for the retrieval of ER‐resident factors as well as ER
export‐associated cargo receptors from the Golgi and the ERGIC back to the ER.

20
The Endoplasmic Reticulum (ER) and the ER‐to‐Golgi intermediate compartment – ERGIC
ER dynamics, morphology and general function
The ER is by far the most extensive organelle within the cell. Rough estimates makes it out
to be a bit more than 10 % of the cell volume [12]. Microscopical analysis of the organelle
reveals that it has a multitude of morphological traits and differences organized in
noticeable regions. It appears to entail features such as sheets, and tubules that form vast
polygonal shapes all interconnected through three‐way junctions.
The extensive membrane network of the ER associates with both the microtubule (MT)
network and actin skeleton to stretch out the organelle into the lace‐like structure that
defines it. Stable attachment and tethering between the ER and MT's are for example
mediated by the ER resident CLIMP63 via its binding to the MT‐bound MAP‐2 protein [13].
Movement of the ER can happen in unison with MT polymerization by the Tip Attachment
Complex (TAC), which connects the ER to the plus‐end of the MT's [14]. The ER can also use
kinesin‐1 connections to slide along acetylated MT's, which also explains the weak effects
MT depolymerization by nocadazole treatment has on ER dynamics [15, 16].
The tubular structure of the ER is maintained by at least two families of membrane proteins,
the eukaryotic membrane‐bound reticulons and the DP1/YOP‐1 protein family [17]. Both
families use a hair‐pin wedging mechanism to distort the membranes. Subsequent
oligomerization of the proteins into arc‐like scaffolds molds the ER layers into tubules [17‐
20]. ER sheets are believed to be generated by the high translational activity on the surface
of the ER. This activity connects multiple translocon‐ribosome complexes across a wide span
of the ER membrane, and as a consequence inhibits reticulon and/or DP1/Yop1 binding [21‐
23].
Finally, the branching of ER tubules into a network is mediated by a class of membrane‐
bound dynamin‐like GTPase proteins named atlastins, in the mammalian system. The
atlastins interact with both the reticulons and DP1/Yop1, and mediate fusion between
different ER tubules [24, 25].

21
EM studies have defined two overall types of ER, the rER and the smooth ER (sER). In these
studies, the bi‐layer of the rER appeared to have a "studded" feature along the outer
surface of extensively stacked sheet‐like cisternae [26, 27]. The "studs" were quickly
identified as membrane‐bound ribosomes. Today we have gained a fundamental knowledge
on how this region plays a vital role in the biogenesis of proteins [28‐30]. The attachment of
ribosomes to the membrane surface occurs through interactions with the Sec61
translocation complex, an ER membrane channel responsible for the transport of nascent
polypeptides into the lumen of the ER [31‐36]. The translocon also controls the insertion of
membrane‐spanning regions into the lipid bi‐layer [37‐39]. Within the ER lumen, newly
formed polypeptides get properly folded with the aid of a variety of chaperones such as BIP,
calnexin, calreticulin and protein disulfide isomerase [40, 41]. If proteins fail to achieve a
native conformation they are targeted for degradation by the ER‐associated degradation
(ERAD) machinery. By this system, misfolded proteins are transported back to the cytosol
where they get tagged with ubiquitin, which destines them for proteolytic degradation by
the 26S proteasome [40, 41].
The major type of ER observed in early EM studies showed a highly convoluted tubular
"unstudded"/smooth structure, implying a different role than protein synthesis [27]. These
tubules have today been recognized as a major site of sterol and steroid synthesis (See
section "Lipids, cholesterol and membrane bi‐layer organization in the cell") [42‐44].
ER exit sites and the ERGIC
As proteins fold and pass ER quality control they are quickly transported towards the Golgi
for further processing, passing through the ERGIC on their way. Specific transitional areas
within the sER termed transitional ER (tER) that are enriched in COPII coated budding
structures in association with vesicular structures, have been recognized as the major hubs
for initiating biosynthetic transport. These structures are what have been defined as ERES.
ERES assemble around an organized center, which is formed by juxtaposition of one or more
tER‐enriched ER cisternae. High levels of budding takes place to fill up the enclosed region
with vesicles that can undergo homotypic fusion. As a consequence, the budded structures
merge to form Vesicular Tubular Clusters (VTC's) (see fig. 1 and 2) [45].

Incubating cells at 10°C has been known to prevent cargo exit and to cause greater
22
abundance of tER structures [46, 47]. Anna Mezzacasa and Ari Helenius have shown that the
cargo in this case is arrested in the ERES. They utilized a useful temperature‐sensitive folding
mutant of the late phase G‐protein from the Vesicular Stomatitis Virus capsid (VSV‐G‐
tsO45), which accumulates in an unfolded state in the ER at 39.5°C. The protein refolds and
exports when switched to 32°C, and can be easily followed throughout the secretory
pathway. They observed that incubation of VSV‐G‐tsO45‐expressing Vero cells at 10°C
caused folded VSV‐G‐tsO45 to accumulate in COPII‐marked ERES, not being able to move to
VTC's or the Golgi [48].
The dynamic VTC's were originally observed as "pre‐Golgi compartments" by Jaakko Saraste
and Esa Kuismanen when they studied the transport kinetics of Semliki Forest Virus (SFV)
membrane glycoproteins. Incubating cells infected with SFV at 15°C they found that SFV
glycoproteins accumulated in defined structures distal to the ER, and identified these
structures as probable ER‐to‐Golgi intermediary stations [49]. These clusters were
subsequently mapped and termed the ER‐Golgi Intermediate Compartment – ERGIC. As
Saraste and Kuismanen presumed, the ERGIC represents a collection of intermediary
stations in the transport pathway between the ER and Golgi where initial post‐ER sorting is
carried out. An interesting feature is that within the ERGIC, COPII‐coated vesicles start to
recruit COPI components, which indicates that ER retrieval is initiated immediately following
budding [50, 51]. Fusion of COPI vesicles that return ER‐specific factors retrieved from the
Golgi, also helps establish and maintain the ERGIC [52].
Figure 2 ‐ ER Exit Complex ‐ Reconstruction of an ER Exit
complex from EM recording. Multiple cisternae (green)
assembled around an organized center. High level of
budding from COPII enriched ERES (see red circle) on the
cisternae fill up the center, where they undergo high level
of homotypic fusion forming vesiculated tubular structures
(VTC's) aka ER‐to‐Golgi intermediate compartments
(ERGIC). (Adapted from Bannykh, S. I. et al (1996)) [45].

23
The ERGIC exists as an organelle that is stabilized through tethering promoted by a specific
hexameric complex named Transport/Trafficking Protein Particle I (TRAPPI). TRAPPI binds to
the COPII component Sec23 and thereby mediates vesicle tethering between individual
COPII vesicles, and between COPII vesicles and the Golgi. It also mediates the homotypic
fusion between the vesicles, and COPII fusion to the ERGIC and the cis‐Golgi. This fusion
event is controlled by the interactions of a set of COPII associated SNARE's (soluble NSF (N‐
ethylmaleimide‐sensitive factor) attachment protein (SNAP) receptors) and their tethers
such as p115 or Rab1 that bridge individual membranes and promote their mixing and
fusion [53‐56].
The ERGIC53 protein
One protein has become synonymous with the ERGIC as a marker of this dynamic organelle,
a mannose‐binding Ca 2+ ‐dependent L‐type lectin of 510 residues named ERGIC53. ERGIC53
is a cargo receptor that in a 1:1 complex with another cargo receptor – the multiple
coagulation factor deficiency protein 2 (MCFD2) – is essential in secretion of two soluble
glycoproteins important in blood clotting, Factor V and VII [57‐59]. Mutations in ERGIC53
have been identified as the cause of a rare bleeding disorder named combined Factor V and
VII deficiency (F5F8D) [58, 60‐62]. When bound to cargo, ERGIC53 associates with the COPII
complex through an FF motif in its cytoplasmic domain [63, 64]. The cargo is released in
response to reduced Ca 2+ as well as acidification at post‐ER compartments prior to arrival at
the cis‐Golgi. The lectin gets retrieved to the ER through a di‐lysine ER‐retrieval signal
recognized by the COPI machinery [65, 66]. Therefore, ERGIC53 appears to be cycling within
the boundaries that make up the ER‐ERGIC interface.
The Golgi apparatus and COPI
A majority of the newly formed polypeptides made in the ER need extensive processing to
mature regardless of their final target destination. A major hub for these post‐translational
processes is the Golgi apparatus. The Golgi is by nature dependent upon a functional COPII
machinery for the delivery of the cargo that needs to be processed. However, the Golgi is
also dependent on COPI for its maintenance as will be evident in this section.

General mammalian Golgi morphology
The mammalian Golgi can be seen by light microscopical methods as a set of stacked
continuous ribbons with perinuclear localization close to, or on top of, the microtubule
24
organizing center (MTOC). The reason for the ribbon morphology is thought to be related to
the polarization of the cell. Trafficking directionality is essential in many polarized cell
functions, such as migration or polarized secretion. Positioning of intact Golgi ribbons helps
the cell to keep an internal orientation, for instance by sensing the apical and basal axis of
the cell or ensuring both directionality and optimal delivery of membrane factors towards
the leading edge of a migrating cell [67‐69]. Inhibiting or perturbing Golgi ribbon formation
does not interrupt global trafficking through the Golgi, but causes major defects in targeting
of polarized secretion and disturbs directional migration during in vitro scratch wounding
assays [70‐72].
The Golgi of the mammalian system is sub‐divided into four sets of compartments; the cis‐,
medial‐, trans‐Golgi cisternae and the Trans Golgi Network (TGN), named according to their
positions in relation to the nucleus. Each compartment contains site‐specific enzymes
involved in the sequential processing of passing cargo. Cargo will generally traverse between
3‐8 cisternae on the way to its final destination, all depending on the type of processing
demands [73].
At the final stage of transport, the cargo passes through the trans‐Golgi and the emanating
reticular membrane network, the TGN [74‐77]. Cargo exits the TGN towards the PM mainly
by vesicular transport, initiated by the activation of the small GTPase from the ADP
ribosylation factor (Arf) family proteins, Arf1. Arf1 binds to clathrin and to a family of
heteromeric Adaptor Proteins (AP‐1, AP‐3 or AP‐4), and γ‐ear containing, Arf‐binding
proteins (GGA's) [78‐86]. The adaptor proteins each recognize a specific collection of signal
motifs in the polypeptide destined for transport, as well as monoubiquitin in the case of the
GGA's [78‐86]. Exit out of the very last TGN cisternae seems to only be mediated through
clathrin coated vesicles, whereas the preceding cisternae are capable of initiating non‐
coated budding and transport [75, 87].

Golgi and cisternal maturation
A long standing model of the Golgi assumed that each individual cisternae was a stable
25
predefined compartment through which secrectory proteins were shuttled for processing.
The shuttling was believed to be maintained by anterograde COPI vesicle trafficking. The
COPI vesicles would specifically sort out and leave Golgi‐resident proteins behind while
trafficking cargo proteins in need of processing. This model provides a good explanation for
the polarity of the organelle and the high concentrations of COPI vesicles observed around
the apparatus [88‐91].
More recent studies have returned to an earlier model, where the Golgi is more likely
maintained by a highly dynamic cisternal maturation process analogous to a conveyor belt
[92, 93]. According to this model, each individual cisternae undergoes a maturation process.
Here, a cis‐Golgi cisternae is assembled by the fusion of COPII vesicles and ERGIC
compartments. The cisternae is then moved through the system from cis‐ through medial‐
and trans‐Golgi, all the way to the TGN, where the cisternae disperses as targeted tubules
and vesicles containing the processed cargo destined for storage or the intended site of
function. In this model, COPI vesicles are assumed to shuttle Golgi‐resident proteins
retrograde from older to younger cisternae [93, 94].
Of the two models, the latter accounts better for the transport of larger molecules, in
particular pro‐collagen, that has been shown not to fit into a conventional COPI and COPII
vesicle. Additionally, no observations to date have been made of pro‐collagen leaving
individual cisternae in specific carriers. Measurements of transport rates show that a
majority of larger cargo, such as pro‐collagen I, moves at the same rate as small cargo
markers such as VSV‐G glycoprotein [95].
COPI
COPI vesicles were initially identified as essential components in maintaining the transport
flow through the Golgi, which explains their primary localization at the cis‐face of the Golgi
[50, 96, 97]. Several similarities have been identified between COPI and COPII. COPI vesicle
formation initiates through Arf1 that gets recruited by a family of oligomerizing cargo
receptor proteins named p24 [98‐102]. The COPI coat assembles in response to Arf1
activation by an Arf Guanosine Exchange Factor (GEF), exchanging a bound GDP to GTP in

the Arf1 [103‐106]. This exchange facilitates the insertion of a myristoylated N‐terminal
26
helix into the lipid bi‐layer of the Golgi surface, thereby tethering Arf1 to the membrane as
well as initiating membrane deformation which starts forming the vesicle bud [107‐112].
The activated Arf1 recruits the preassembled COPI complex consisting of the following 7
subunits: α‐, β‐, β'‐, γ‐, δ‐, ε‐ and ζ‐COP [113‐119].
Recent advances in the crystallization of the COPI complex have indicated that the coat
most likely assembles into a quarternary structure similar to the well‐known triskelion of
clathrin, but with a tertiary structure subunit that resembles the basic COPII Sec13/31
associations (see fig. 3) [120]. A very recent study has shown that Arf1 binds both to the γζ‐
COP and βδ‐COP, meaning that each COPI coatomer associates with the lipid membrane
through two Arf1 molecules [121].
Figure 3 ‐ Comparison of COPI, COPII cage and the clathrin
triskelion – Top: The crystal structure of assembled COPI complex,
with solenoid arms curving outwards from an assembled hinge
made out of β‐COP β‐propeller domains. Bottom: Graphic
comparison of COPII, COPI and clathrin cage structures. Notice the
similarities between the sub‐unit architecture of COPII and COPI
with β‐propeller domains forming the hinge of each cage vertice.
Also notice the curvature of the assembled COPI vertices that
have apparent similarities to the known structure of the clathrin
triskellion represented to the far right (Adapted from Lee, C. and
Goldberg, J. (2010)) [120].

COPI cargo loading
27
The loading of cargo into the COPI vesicles is controlled either by direct interaction of cargo
transport motifs with the coatomers ‐ as observed for membrane‐bound cargo ‐ or through
a loading machinery that helps retrieving soluble cargo to the ER.
Classic examples of transport motifs for membrane‐bound cargo are the dilysine motifs
KKXX and KXKXX, found in a wide variety of ER proteins. Retrieval is mediated by direct
interactions with the α‐ and β'‐COP subunits [97, 122‐128]. Lumenal cargo, on the other
hand, carry a KDEL or KDEL‐like sequence that directs their binding to KDEL‐recognizing
receptors (KDELR) located within the cis‐Golgi. This re‐directs the cargo back to the ER by
association of KDELR's with COPI vesicles [129‐131]. The KDELR's dissociate from their cargo
in response to the pH change from acidic to neutral observed between the cis‐Golgi and the
ER, and recycles to the early Golgi for additional rounds of transport [132].
Uncoating of COPI vesicles, a necessary step prior to fusion with the target membrane, is
triggered and controlled by Arf GTPase Activating Proteins (ArfGAPs), which catalyze the
hydrolysis of the Arf‐bound GTP [133‐135].
It is important to note that COPI and COPII activities are coupled. Blocking COPI dependent
retrograde transport causes inhibition of COPII‐mediated anterograde transport. Because
COPI is largely implicated in retrograde trafficking, transport of novel proteins is apparently
highly dependent upon the efficient return of the Golgi targeting factors that escorted the
previous batch of the COPII‐associated cargo [50].
COPII
COPII was initially discovered using yeast genetics. Temperature‐sensitive mutations within
a specific set of genes were shown by Peter Novick, Charles Field and Randy Schekman to
inhibit the transport of marker enzymes [136]. Protein production was observed to be still
ongoing, while vesicular clusters or expanded ER membranes accumulated. Subsequently,
the identified genes were termed SEC (secretory deficient) [136‐139].

The screen revealed a vast and intricate network of participating proteins. Among these,
seven participate in the budding on the ER: Sec12, Sar1, Sec23, Sec24, Sec13, Sec 31 and
28
Sec16. These are the essential components of the COPII complex. These proteins have been
shown to co‐operate in an ordered fashion to both ensure vesicle formation as well as
selection and packaging of cargo into budding vesicles (see fig. 4) [140].
Figure 4 ‐ COPII recruitment and budding‐ Sequence in the formation of COPII vesicles. The Sec12 GEF recruits and tethers
Sar1 to the ER membrane by exchanging a bound GDP to GTP. In turn, Sar1 recruits the inner layer of the COPII coat
consisting of the GAP Sec23 and the cargo receptor Sec24. Next the Sec13/Sec31 gets recruited, stabilizing the cage
structure and catalyzing the membrane constriction that leads to release of the vesicle (Adapted from Sato, K. (2004)
[143]).
Sec12
Sec12 resides predominantly on the cytoplasmic surface of the ER [141‐143], tethered to
the membrane by a C‐terminal domain [144]. Sec12 recruits and activates a small GTPase
from the Ras superfamily, Sar1, which belongs to the Arf family [145‐148]. The recruitment
of Sar1 by Sec12 causes a conformational change in Sar1 that mediates its tethering to the
ER membrane [149‐152]. This recruitment initiates the budding process and formation of a
COPII cargo vesicle (see fig. 4).
The cytosolic domain of S. cerevisiae Sec12 has recently been crystallized. The protein folds
into a seven blade β‐propeller. An extended loop dubbed the "K‐loop" projects upward from
the first propeller blade. This loop has been shown to bind K + and has been identified as

29
important in the Sec12‐Sar1 interaction as it enhances Sec12 GEF activity [153]. Sec12 and
Sar1 work together to initiate COPII vesicle formation. Currently, Sec12 is the only GEF
known to activate Sar1 [154].
Sar1
Sar1 was initially identified as a suppressor of the temperature‐sensitive Sec12 mutant
[145]. Sar1 exists in two isoforms in mammalian cells, Sar1A and Sar1B. The major functional
difference between these isoforms is that Sar1B appears to be essential in the transport of
chylomicrons from the ER [155, 156].
Similar to most small GTP'ases, Sar1 contains a structural core consisting mainly of a
nucleotide binding pocket where a Mg 2+ ion aids in holding the GDP molecule. Moreover, a
threonine at position 39 (T39) within the pocket is essential for GTP binding (see fig. 5)
[152]. A mutation substituting the threonine with an asparagine (T39N) interferes with
interactions necessary for Sec12 to induce the essential nucleotide exchange, and renders
the protein locked in a dominant negative GDP‐bound state [152].
Sar1 contains two switch regions (I and II) positioned on either side of the nucleotide
binding pocket [152]. These regions change their conformations in response to the bound
nucleotide. The switch II region contains a histidine at position 79 that is essential for the
GTP hydrolysis reaction. Changing this residue to a glycine (H79G) locks the protein in a
constitutively active GTP‐bound state, inhibiting the disassembly of the formed complex and
vesicle fission (see fig. 5) [146, 147, 152, 157, 158].
Figure 5 ‐ Sar1A – The Sar1A structure seen from three angles (from left to right), 1) back with the nucleotide pocket
turned away from view, 2) front with the nucleotide binding pocket turned towards the viewer and 3) sideways. The white
arrows shows a bound GDP in the nucleotide pocket of the protein, and the red arrows show the switch regions (modeled
from PDB accession # 1F6B ‐ Sar1A bound with GDP‐ using 3D‐molecule viewer (Invitrogen)).

A distinct feature of Sar1 is that it does not contain any prenyl‐lipid modifications for
membrane binding and tethering at the N‐terminus, a feature found in the close relatives
30
Arf1 and Rab. Instead, the protein utilizes an extended amphipathic N‐terminus that inserts
into the lipid bi‐layer, when the protein is in a GTP bound state, and thereby tethers the
protein to the membrane [149‐152]. The insertion of the N‐terminus furthermore extends
the surface of the outer leaflet. This starts a nucleation process, where additional Sar1 gets
recruited and organizes into a helical protofilament‐like scaffold leading to membrane
tubulation [150, 159]. Accommodation of the area expansion of the outer leaflet induces
elastic stress on the inner leaflet, which in turn accommodates by causing a local change in
the membrane shape such as forming a tubule [160]. Additional organized local clustering of
the Sar1 N‐terminus results in the tubule further constricting to a 'beads‐on‐a‐string" like
morphology. This is believed to cause sufficient perturbation of the inner leaflet
organization for the hydrophobic interior of the membrane to become exposed. This causes
the interior to collapse and thereby drive fission [149, 150, 152, 159, 160]. GTP hydrolysis of
Sar1‐GTP promotes the final steps of fission [150, 159].
The activation of Sar1 by GTP loading is essential for the recruitment of the inner layer COPII
components, Sec23/Sec24 (see below). Hydrolysis of Sar1‐bound GTP results in rapid
disassembly of the recruited COPII coat [45, 157]. The recruited coat itself induces the
hydrolysis of the Sar1‐bound GTP [161]. All this implies that a very fine balance is
maintained between COPII recruitment/coating, the un‐coating of the bud and
constriction/fission. Here Sar1 is intrinsically involved in controlling the retention time of the
coat on the bud, but is also itself controlled by interactions with the coat to induce a
productive vesiculation.
Several lines of evidence suggest that factors other than Sar1 may influence coat residency
on the membrane. For instance, changes in the local lipid environment can promote
budding and fission (see the section on Membranes and lipids) [159, 162]. These changes
can be promoted or responded to by accessory proteins, such as Tango1 in concert with
Sedlin [163, 164]. On the bud, Tango1 and Sedlin control and extend the coat life‐time and
the Sar1 cycle, respectively, to ensure loading of large cargo, i.e. pro‐collagen [163, 164].
Another example is Sed4 in yeast that promotes GTP hydrolysis in Sar1 when the coat
associates to the bud site without cargo [165, 166].

Sec23 and Sec24
The Sec23/Sec24 complex has three major roles during COPII vesicle formation; 1) Cargo
31
binding (which particularly involves Sec24) to ensure proper packaging of cargo protein into
budding COPII vesicles; 2) membrane lipid binding, which provides the interactions between
the budding lipid membrane and the forming COPII complex, and 3) GAP activity, whereby it
provides a mechanism to control the life‐time of the complex at the bud site and also
control Sar1 membrane interaction.
Activation of Sar1 leads to recruitment of the inner layer of the COPII complex through
direct interaction with a conserved region on the surface of Sec23 (see fig. 4 and 6) [151,
167, 168]. X‐ray structures of both the Sar1/Sec23 and Sec23/Sec24 complexes have
revealed that they are highly refined for the interaction with a curved surface of a vesicle.
The Sec23/Sec24 complex forms an intricate bow‐like structure in cooperation with Sar1,
with an inside curvature that fits a standard 60 nm vesicle (see fig. 6) [151, 169, 170]. A
striking feature of Sec23 and Sec24 is that they fold into virtually homologous structures,
even though they only share about 14 % sequence similarity. Both proteins fold into a
twisted triangular shape. Sec23, in contrast to Sec24, furthermore encompasses contact
sites to interact with Sar1 (see fig. 6) [151, 171].
Figure 6 ‐ The GAP Sec23 (yellow) bound to Sar1 (red) in complex with the cargo receptor Sec24 (green) seen from three
angles ‐ Angle b shows the complex when bound to the vesicle surface, with the Sar1 N‐terminus inserted into the
membrane leaflet (red arrow), and the concave surface of Sec23/Sec24 associating with the curved membrane surface.
(Adapted from Bi, X. et al (2002) [151]).

32
Two specific sites in Sec24 have been identified that are involved in cargo recognition, the A‐
site and the B‐site. The A‐site forms a pocket located on the periphery of the membrane‐
proximal surface of Sec24 [172]. The B‐site constitutes a shallow groove on the periphery of
the membrane‐interaction surface of Sec24. The B‐site is known to associate with the DxE
export motif of VSV‐G [172‐174]. A third cargo interaction site, named the C‐site, was
identified through a point mutation that influenced COPII interactions with the SNARE
protein Sec22 [175]. Co‐crystalization of Sec23/Sec24 with Sec22 revealed that the
interaction site was located in the Sec23/Sec24 interaction groove, and that Sec22 is
recognized by Sec23 and Sec24 not by a conserved motif, but rather a conformational
epitope [173, 175].
A catalytic arginine at position 722 (R722) from Sec23 interacts with Sar1 by inserting into
the nucleotide binding pocket of Sar1. The arginine guanidinium group interacts with the
phosphates of the nucleotide, and helps neutralize negative charges formed by the
transition state during the hydrolysis (see fig. 7B) [151].
A recent study has discovered a novel mutant of yeast Sec24, called m11, containing two
point mutations (E504A and D505A) on a surface loop flanking the A‐site [176]. These
mutations perturb Sec24p association with the scaffolding protein Sec16p. As a
consequence, Sec16p mediated inhibition of the COPII promoted Sar1‐GTP hydrolysis is
inhibited, and a decrease in packaging efficiency as well as an increase in small vesicle
formation can be detected [176]. These observations add to the function of Sec24 beyond
just cargo binding and loading.
Sec13 and Sec31
The association of the inner Sec23/24 layer with Sar1 signals for the recruitment of the
outer layer of the COPII complex that consists of a heterotetramer formed by the two
proteins Sec13 and Sec31. The Sec13/31 heterotetramer forms a "ball‐capped" rod
consisting of a central α‐solenoid region comprised from the two C‐termini of Sec31, which
shape the rod [177]. The "ball‐caps", at each end of the rod, are assembled through a
characteristic evolutionarily conserved crown, trunk and tail motif named Ancestral
Coatomer Element 1 (ACE1) within Sec31 (see fig. 7A) [177, 178].

The ball cap constitutes the crown in ACE1 and the α‐solenoid forms the trunk. The tail
33
associates with Sec13 forming a second ball like structure just beneath the crown (see fig.
7A). The crowns of each individual heterotetramer associate with each other in an off‐edge
assembly, forming the vertices of the COPII cage (see fig. 7A and fig. 8) [179, 180].
Sec31 contacts Sec23, but not Sec24, through a 50‐residue fragment from an unstructured
region. This Sec31 region binds across the surface of Sec23 and extends a short N‐terminal
element into the nucleotide‐binding pocket of Sar1 (see fig. 7B) [181]. A tryptophan in
position 922 (W922) and an asparagine in position 923 (N923) in the Sec31 fragment
interact with the catalytic H79 of Sar1, forming a lid to the pocket cavity of Sar1. This orients
H79 for catalysis, and also accelerates the Sec23‐promoted catalysis up to ten‐fold [151,
161, 181].
Figure 7 ‐ A) Sec13 and Sec31, B) Sec23 and Sar1A
associated with a Sec31A fragment ‐ A)
Heterotetramer complex of Sec13 (red and orange)
and Sec31 (dark green and light green). The C‐termini
of Sec31 associate to form the α‐solenoid rod (trunk)
with a crown as a ball cap. Sec13 (red and orange)
binds to a tail, protruding from the trunk of Sec31,
forming a second ball beneath the crown (Adapted
from Fath, S. et al (2007) [177]). B) Sec23 (yellow)
associating with Sar1A (red) promoting the hydrolysis
of the bound nucleotide. Also seen in turquoise/blue
is the active Sec31A fragment binding across the
surface of Sec23 donating a tryptophan (W922) and
an asparagine (N923) that catalyze nucleotide
hydrolysis in Sar1A (Adapted from Bi, X et al (2007)
[181]).
There is some debate regarding the flexibility of the assembled cage. Early cryo‐EM data of
the self‐assembled Sec13/31 outer cage components revealed that they have a propensity
for associating into cuboctahedral cage‐like particles with an average diameter of app. 600 Å
(60 nm) (See Fig. 8) [182]. In this configuration, large cargo such as chylomicrons and pro‐

collagen cannot be accommodated. Speculations were made over the possibility that the
cage could expand by addition of more vertices, creating larger shapes, e.g. octahedrons,
34
icosidodecahedrons or small rhombicosidodecahedrons [182]. Later cryo‐EM experiments
were able to identify icosidodecahedral structures when Sec13/31 was assembled with
Sec23/24. In these studies, it was plain to see that the majority of the accommodations
occur at the vertex interfaces extending the angles between the assembled heterotetramers
[177, 180, 183].
Budding from artificial liposomes has been achieved by mixing just Sar1 with the inner and
outer coat components and GMP‐PNP (a non‐hydrolysable analog of GTP), although the
findings did show that liposomes needed to be composed of acidic lipids to induce
recruitment of the coatomers [184]. The electrostatic interactions between the negatively
charged lipids of the liposome membrane and positive charges within the membrane facing
curvature of the Sec23/24 heterodimer are believed to help stabilize the Sar1‐Sec23
association and overall COPII assembly [151, 169, 170, 184, 185].
Figure 8 – Cuboctahedral COPII cage on vesicle ‐ A)
Assembly of the COPII cage on a vesicle, with the Sec31
crowns forming the hinged vertices of the cuboctahedral
cage and the α‐solenoid stretching across the vesicle
surface. Sec23 (purple)/Sec24 (blue) bound with Sar1
(red) is seen below a vertice on the very left of the cage
(Adapted from Fath, S. et al (2007) [177]).
The tight association of the two coat layers with lipids, cargo and accessory proteins causes
the lattice to linger upon the bud/vesicle membrane after the hydrolysis of GTP in Sar1
[186]. Accessory proteins are non‐COPII factors that have recently been identified to
associate with Sec13/31. One such accessory protein is the apoptosis linked gene 2 (Alg‐2)
that has been shown to modulate the COPII disassembly kinetics at the ERES in response
increases in Ca 2+ flux by binding to a proline‐rich region within Sec31 [187‐189]. Also signal‐

35
transducing adaptor molecules (STAM's), which are involved in growth factor and cytokine
signaling as well as receptor degradation, have been identified to associate with Sec31A in
co‐immunoprecipitation experiments [190]. Overexpression of STAM2 caused a decreased
intensity of the Sec31 signal at ERES and cytocolic re‐distribution of Sec24. The recruitment
of STAM2 to ERES was further shown to be Sar1 dependent and not influenced by the
Sec31A expression levels. Both depletion and over‐expression of STAM2 caused inhibition of
VSV‐G transport [190].
Cargo loading and ER export motifs
Cargo loading has been shown to be mediated by interactions with Sar1 as well as the inner
coatomer layer, in particular with the Sec24 subunit, during assembly of the ERES [168, 191,
192]. Sec24 specifically binds to the ER export motifs in the cargo molecule C‐terminus [157,
175, 191, 193‐197]. It is interesting to notice that Sec24 has several orthologoues in both
the yeast and the mammalian system that each respond to different cargo motifs [198‐200].
On the other hand, orthologoues of Sec12, Sar1, Sec23, Sec13 and Sec31 have not been
found in the yeast system but are present in the mammalian system [198, 201, 202].
Different ER export motifs for transmembrane cargo have been recognized to bind to e.g.
Sec23/24. A well‐characterized export motif comprise of a di‐acidic cluster, such as the
YxExD/DxE seen in the carboxy tail of VSV‐G [174, 203, 204]. Other known export motifs
involve hydrophobic or aromatic amino acids at the C‐terminus, e.g. two phenylalanine (FF)
residues of the cargo export receptor ERGIC53 or p23/24 family proteins [124, 197, 199], or
a single valine in as seen with CD8 [205].
A different export mechanism has been identified for Sec22, where protein folding creates a
specific conformational epitope that is recognized as a transport signal by both Sec23 and
Sec24 [173]. This also suggests that the COPII machinery may act as a component of the
quality control system in the ER, since only proper folding of a protein presents a useful ER
exit signal [173].
Soluble cargo has been proposed to package into vesicles indiscriminately by a "bulk flow"
mechanism. Unintentionally transported proteins, such as ER‐resident proteins, would then
be retrieved by an intrinsic sorting signal, e.g. KDEL [206]. The bulk flow mechanism was
adapted from observations in plants, where cargo was observed to export via a default

36
pathway [207]. Studies with pro‐α‐factor in yeast seemed to imply that a receptor‐mediated
transport was also present in ER export [168]. The responsible cargo receptor was later
identified as ERv29p, a membrane bound protein known to interact with COPII [208‐210].
Other cargo receptors such as the previously mentioned ERGIC53 and MCDF2 have later
also been identified [57‐59]. Whether a bulk‐flow mechanism is present needs still to be
determined [211, 212].
COPII mutations and physiological effects
The important cellular function of COPII‐mediated transport is reflected by a number of
serious diseases caused by COPII dysfunction. At the same time, research into the
underlying molecular mechanisms of these diseases illustrates important principles of COPII
assembly and trafficking.
Of the genetic diseases that have been identified as correlated with defects in the COPII
machinery, many are related to the deposition of connective tissue and in particular with
the secretion of collagen from cells [213, 214]. A particularly interesting mutation in a sub‐
type of Sec23, Sec23A, causing a substitution of a phenylalanine to a leucine at position 382
(F382L), manifests in humans as bone diseases and problems with closure of the fontanel
causing cranio‐lenticulo‐sutural dysplasia. The mutant Sec23A is still capable of associating
with Sec24 and Sar1 and initiating the budding process, but Sec13/31 does not get recruited
to the budding site. Since the mutation lies within a part of Sec23 that has been identified to
associate with the catalytic region of Sec31, this mutation is believed to inhibit productive
binding of Sec31 to Sec23, and subsequent Sec31 association with the catalytic site within
the Sar1 sub‐type, Sar1B [181, 215, 216].
Further evidence for the necessity of Sec23 and Sec31 association for pro‐collagen transport
has been found in a novel Sec23 point mutation, changing a methionine at position 702 to a
valine (M702V) right next to the aforementioned F328L in the fully folded protein [217]. This
mutation still retains Sec31 recruitment ability, but the association causes accelerated
activation of Sar1B GTP hydrolysis, and manifests itself by defects in pro‐collagen transport.
The mutation is still able to maintain transport of smaller molecules e.g. ERGIC53, Sec22 and
amyloid precursor protein. This suggests that a prolonged association of the four COPII

37
coatomer subunits is essential in promoting a productive packaging of large cargo molecules
such as pro‐collagen [217].
Sar1B mutations have been identified in people with lipid processing defects, such as
Anderson's disease or chylomicron disease that presents itself in infants as a failure to thrive
as well as chronic diarrhea. The reason for these symptoms has been identified as a failure
by enterocytes to secrete chylomicrons into the lymph. These lipids particles are instead
retained within the ER causing low lipid levels in the plasma, and this in turn causes a
detrimental decrease in available fat‐soluble vitamins that usually manifests in neurological
impairments [155, 218]. Similar lipid disorders are also associated with mutations in Sec24C
that prevent pre‐chylomicron vesicles to dock with the Golgi [156, 219]. Mutations in
Sec23B have been identified in defects associated with erythrocyte maturation, causing
congenital dyserythropoietic anemia [220]. Mutations in Sec24B have been shown to affect
planar cell polarity, causing major developmental defects in mice such as craniorachischisis,
neural tube closure defects, disturbed cochlea development, to name a few [221].
Recent knock‐outs in mice of both Sec23A and B as well as Sec24D have turned out to be
embryonic lethal. Knock out Sec23A animals die mid‐embryogenesis and Sec23B knock‐outs
showed massive pancreatic degenerations and perinatal lethality. Sec24D knock‐outs were
aborted prior either to the blastocyte stage or between E10.5 and E18.5 with none surviving
to the latter. The varying effects where ascribed to differences in intron insertions of the
gene trap [222, 223].
The assembly of both COPI vesicles and of COPII vesicles has been shown to be very
dependent upon the local membrane environment. Several factors governing recruitment of
the components as well as shaping the membrane are controlled by reactions that target
and modify the individual lipids, which make up the membranes. These modulations will be
described in the following section.

Membranes and lipid biogenesis
38
In 1972 S.J. Singer and Garth L. Nicolson proposed a view of the biological membrane as "a
fluid mosaic" where proteins floated and interacted in a two‐dimensional sea of lipids, and
the lipids acted more or less as "solvent" where "(…) a small fraction of the lipids may
interact specifically with the membrane proteins." [224]. Today we know that lipids and
membranes are far more dynamic, and exert far more influence on proteins and the
functions of the cell than Singer and Nicolson envisioned in their model.
Membranes are an integral part of the cell, where they compartmentalize several vital
processing centers such as the nucleus, Golgi, the mitochondria and the ER. Cellular
membranes are composed by a large variety of lipids. Most membrane lipids consist of two
hydrophobic acyl tails and a hydrophilic head group. The predominant head groups found in
phospholipids are usually derived from either serine, choline, ethanolamine or the sugar
inositol. The lipids are furthermore classified by said head group derivate (see fig. 9) [225].
The acyl tails vary in length, usually 14 to 24 carbons, with one of the chains being poly‐
unsaturated and the other either mono‐unsaturated or saturated [225, 226]. This high
degree of variability means that the cell uses more than 1000 different types of lipids in the
assembly of membranes [227].
The majority of phospholipid biogenesis occurs upon the cytosolic leaflet of the ER
membrane [228, 229]. The ER furthermore synthesizes Ceramide (Cer), the precursor for
most of the glycolipids and sphingomyelin (SM) that are mainly produced on the Golgi [230,
231]. The Golgi also has the capability of producing phosphatidylcholine (PC) and
phosphatidylethanolamine (PE) by head group substitution (see fig. 9) [232, 233].
Phosphatidic Acid (PA) is synthesized on the cytosolic leaflet of the ER by acetylation of
glycerol‐3‐phosphate (G3P) [225, 234‐237]. The formed PA is subsequently
dephosphorylated into diacylglycerol, and serves as a precursor for the formation of PC, PE,
PS and triacylglycerols [225]. Alternatively, the PA is converted into CDP‐diacylglycerol (CDP‐
DAG) [225]. CDP‐DAG gets converted to either cardiolipins, phosphatidylglycerol (PG) or to
phosphatidylinositol (PI) [225, 234‐237]. The latter, PI, is produced through the ER and PM
resident PI synthase, which substitutes the CDP molecule of the CDP‐DAG with a myo‐

inositol molecule releasing a CMP in the process [237‐239]. PI and its phosphorylated
derivatives are important for signal transduction and in initiating vesicle trafficking in the
cell [240‐242].
39
The most abundant phospholipid within the cell is PC, which makes up for app. 50 % of the
total phospholipid mass of a eukaryotic cell. Second most abundant is PE, which makes up
for app. 25 % of the total lipid mass in a eukaryotic cell [232].
Lipid transport
The lipid composition varies greatly between the various organelle membranes of the cell
[226, 227]. How the cell distributes the various lipids from their site of synthesis has still not
been mapped properly [243]. Within membranes, lipid composition is managed by
scramblases, flippases and floppases that are capable of translocating lipids between the
two leaflets [244‐248]. Inter‐organellar lipid exchange is presumed to occur either via vesicle
trafficking, or phospholipid exchange proteins that are capable of extracting lipids out of
one membrane, shielding it from the surrounding aqueous environment while delivering the
lipids to target membranes [225, 249‐252].
Whether specific lipid sorting happens during the formation of transport vesicles on the ER
is still not fully known. Studies with SM and glycerophospholipids imply that their
movement out of the ER is maintained by bulk flow into COPII vesicles [253, 254], whereas
inhibiting Golgi traffic or disrupting the cytoskeletal railing system appear to have no
influence on the distribution of PE or PI to the PM [255‐257].
Figure 9 – Phospholipids and their
head groups – The four main
phospholipids and their associated
head groups. Note that carbon
position 1 in the inositol sugar is
attached to phosphate of the
diacyl‐phosphoglycerol (Adapted
from
http://resources.jorum.ac.uk/xmlui
/bitstream/handle/123456789/138
08/page29.htm).

Lipid exchange may occur via contact sites between the ER and the recipient organelle
40
membrane. The lipid exchange occurs either by diffusion across the short cytosolic span or
the lipids may be carried across via specific lipid transfer proteins. One example is the
recruitment of Cer transfer protein (CERT) and Vesicle Associated membrane Protein (VAP‐A
and VAP‐B), which promote shuttling of Cer from the ER to the Golgi by connecting a
juxtaposed ER region to a trans‐Golgi cisternae [258‐261]. Similar examples are found
connecting the ER to the mitochondria or the PM, where fractionation of these regions have
shown them enriched in PS synthase [262‐268].
Cholesterol and membrane fluidity
The fluidity and the shape of the lipid membrane can be varied in response to the
heterogeneous features of the composing lipids. Varying lengths and levels of saturation in
the acyl chains can cause varied alignment within in the bi‐layer [269‐272]. The easier the
acyl chains are able to align, the higher degree of order is achieved, to a point where the bi‐
layer becomes rigid and gel like. This is termed the membrane's solid state (so) [273].
Disordered acyl chain alignment tend to give the membranes a morphology resembling
crystalline liquid, and is termed liquid disordered phase (Ld) [273]. By incorporation of
sterols, and in particular cholesterol, into the bi‐layer, the membranes can achieve a
transitional state between gel and crystalline state, termed liquid ordered phase (Lo)[273].
The cholesterol thereby aids in organizing and aligning the acyl chains, mixing so‐ with Ld ‐
preferring lipids and maintaining them in an Lo phase [274, 275]. It is believed that the
decrease in line tension in the boundary between the Lo and Ld favors the Ld phase to bulge
which thereby supports vesiculation [276, 277].
Cells acquire cholesterol through two pathways: either by a complicated synthesis pathway,
involving more than 30 different enzymes, that condensate acetyl‐Coenzyme A over several
turns to yield cholesterol [278, 279]; or the cell acquires the cholesterol by receptor‐
mediated endocytic uptake of low‐density lipoprotein following sorting out through the
lysosomes [280].
Cholesterol can be fractionated from a variety of eukaryotic cells in detergent resistant
membrane (DRM) fractions that are also enriched in sphingolipids [274, 281]. Suggestions
have been made that the DRM's exist as insoluble rafts on the membrane used to tether,

sort and transport associated proteins, i.e. glycosylphosphatidylinositol (GPI)‐anchored
proteins from the Golgi to the PM [282, 283].
41
Although sterols and cholesterol are synthesized on the ER membrane, they only constitute
a few percent of the ER membrane's total lipids [284, 285]. Still, they play an important role
in cargo packaging at the ERES. Cells cultivated in lipoprotein depleted serum and
subsequently exposed to 2‐hydroxypropyl‐β‐cyclodextrin, which causes extraction of
cholesterol from the cell, showed a significant delay in VSV‐G‐ts‐O45‐YFP transport from the
ER to the Golgi [286]. FRAP of the ERES, and in particular the COPII component Sec23,
revealed that the turn‐over of Sec23 had increased in the cholesterol‐depleted cells,
suggesting an inhibition in COPII function as a consequence of the treatment [286].
However, the direct mechanistic role of cholesterol in packaging has yet to be deduced. It
has been suggested that initial raft formation and raft‐induced protein sorting may occur
already at the ERES, and depletion of cholesterol would therefore cause a sorting delay
[286‐289]. It has also been implied that dynamic cholesterol micro‐domains directs Sar1
activity at ERES, and furthermore decreases the membrane elasticity at the bud site causing
lipid packaging defects that promote fission and subsequent release of the vesicle [159,
289].
Recruitment of yeast Sar1p to synthetic liposomes has shown an increased nucleotide‐
independent binding of Sar1p when lysophospholipids and oleic acid were added. The same
study also showed a difference in Sar1p binding ability on synthetic liposomes with
variations in the phospholipid saturation levels. Both these experiments showed the need
for a certain level of membrane fluidity to exist to support efficient Sar1p binding [184].
Lipids and membrane curvature
The shape of the actual lipid molecule is used by the cell to promote or discourage the
formation of domains and vesicles [290]. The influence of the area occupied by the head
group in comparison to the volume occupied by the acyl chains of the lipid affects the
internal organization of the membrane, and can cause the membrane to curve
spontaneously [290]. Lipids can be divided into three different shapes: cylindrical, conical
and inverted cones, with each shape promoting a different type of organization, i.e.
membrane bi‐layer, negative curvature and positive curvature (see fig. 10) [269, 290].

The cell is able to control membrane curvature by changing the lipid composition globally
42
and locally [291]. This can be done through; 1) adding or removing acyl chains of lipids, e.g.
PA conversion to LPA by phospholipase A2 removing an acyl chain, or LPA conversion to PA
by lysophosphatidic acyl transferase (LPAT) [291, 292]; 2) substituting or modifying the lipid
head group thereby changing the area occupied by the head group [293]; 3) flippase‐
mediated re‐distribution of the lipids between the bi‐layer leaflets [294‐296].
Membrane curvature can also be induced and controlled by protein interactions and
insertions [291]. Microtubules and cytoskeletal elements can use bundles to protrude and
push local areas of a membrane, or pull out membrane tubules using kinesin motors running
along microtubule tracks [291, 297, 298]. Scaffolding can be mediated by binding of a rigid
protein structure with intrinsic curvature to a membrane surface, which causes the
membrane to bend, as seen with the BAR domain of dynamin [299‐301]. Scaffolding also
occurs as stabilization of induced curvature by coat protein polymerization, as seen with
clathrin, COPI and COPII coats [180, 299, 302]. Local spontaneous curvature can be induced
by insertions of amphipathic helices between the polar head groups of a leaflet in a bi‐layer.
The induced elastic stress on the inner leaflet by the local expansion of the outer leaflet
promotes a change in the membrane shape. This can be seen in protein‐membrane
interactions of epsin and, as previously mentioned, the small GTPases such as Arf1 and Sar1
[111, 149, 159, 303].
Figure 10 – Lipid geometry and spontaneous
curvature. Graphical description of lipid geometry.
Variations in head group size compared to the volume
occupied by the acyl chains. PC and PI, have a
cylindrical shape and spontaneously form bi‐layers.
Conically shaped lipids such as PE, with smaller head
groups compared to the volume occupied by the acyl
chains, have a tendency to spontaneously form
membrane layers with negative curvature. Inverted
cones, where the head group is substantially larger
than the volume occupied by the acyl chain(s) e.g.
LPA, are more prone to associate in membrane layers
with positive curvature, and easily form micelles.
Membrane curvature at ERES is primarily influenced by the actions of Sar1 and the insertion
of its N‐terminal amphipatic helix [149, 150, 159]. In addition, several indications of lipid

organization and modification have been found to influence the stability of ERES. The
43
phorbol ester analogs calphostin C and phorbol 12‐myristate 13‐acetate have been shown
to influence the export of VSV‐G from the ER [304]. Phorbol esters are known to mimic DAG
[305]. As DAG has a relatively small head group it can potentially induce a negative
membrane curvature [290, 306]. It was shown that calphostin C inhibited the export of VSV‐
G from the ER, and that PMA had an opposite effect, stimulating VSV‐G ER export [304]. It
should be noted though, that DAG also serves as signaling molecule, and recruitment of
additional membrane modulating factors could not be ruled out [304].
PA is known to induce negative curvature in membrane bi‐layers [290, 307], and plays an
important role in the assembly and stabilization of COPII at the ERES [308‐310].
Overexpression of diacylglycerol kinase δ (DAGKδ), which phosphorylates DAG into PA, has
also been shown to re‐distribute Golgi markers and to inhibit ER export of VSV‐G [309].
Ethanol‐induced inhibition of phospholipase D (PLD), a protein that catalyzes the formation
of PA, has been observed to inhibit VSV‐G exit from the ER [308]. PLD activity is stimulated
on the ER in response to Sar1A activation, and PLD activity influences Sar1‐promoted
tubulation at ERES [310]. The study was able to show that PA enhanced Sar1A‐dependent
recruitment of the Sec23/24. These studies suggest that introduction of PA during ERES
assembly may promote negative membrane curvature and thus support tubule and vesicle
formation [308‐310].
The presence of the small LPAT antagonist called CI‐976 during temperature‐induced
transport of ts‐O45‐VSV‐G caused a general inhibition of ER export which also blocked the
viral protein from budding at ERES, concentrating VSV‐G at ERES foci. Interestingly, Sar1‐
induced tubule formation during VSV‐G transport in semi‐intact cells was enhanced in the
presence of CI‐976 [311]. These observations imply that a remodeling of the lipid bi‐layer,
from positive to negative curvature, during a late stage of the vesicle formation at the ER is
necessary [290, 307, 311].
PI and phosphorylated PI (PIP): their role in signaling
More and more evidence now support that lipids play a vital role in cell signaling. These lipid
signaling events has proven to be essential for the productive formation of ER export
vesicles. It has also been found that most of the lipids are utilized in relaying signals within

44
the cell. Examples include PA's participation in Ras signaling activation [312], and PS cellular
externalization during apoptosis, which signals for macrophage engulfment and clearing of
the apoptotic cell without causing inflammation in the surrounding cells [313].
A majority of transport initiation in the cell is tightly connected to a particular modification
of PI, namely phosphorylation. Phosphorylations of PI change the head group area and
thereby the geometry of the PI lipid towards an inverted cone shape. Thereby, they may
induce positive curvature [314, 315]. However, due to their ability to become rapidly
phosphorylated at the 3', 4' and/or 5' position of the inositol ring, PI and its phosphorylated
derivatives have been mainly identified as involved in lipid mediated signaling [316].
For instance, PM PI(4,5)P2 is used to mediate targeting, docking and priming of exocytic
vesicles to the PM, thereby influencing vesicle fusion and cargo release [317, 318].
PIP(4,5)P2 also recruits several cytosolic clathrin adaptors, e.g. AP‐2, AP180 and epsin, as
well as the clathrin triskelion during endocytosis [319‐326]. Early endosomes are enriched in
PI(3)P, and this lipid is also essential in the biogenesis of MVB's and in the formation as well
as the maturation of autophagosomes [318, 321, 327‐329]. PI(5)P has been identified in
controlling p53 mediated DNA damage repair, and in membrane trafficking from endosomes
to the PM [330]. And finally, PI(3,4,5)P3 has been found as a transient signal connected to
cell proliferation, metabolism and apoptosis [331].
PI typically represents less than 15 % of the total amount of phospholipids found in the
eukaryotic cell, and as low as 1 % in total lipid by weight in erythrocytes [324, 332]. PI serves
mainly as substrate for the PIP's during signaling. PIP's are less abundant than PI, between a
10‐ to a 100‐fold, with the majority comprising of PI(4)P and PI(4,5)P2 [324, 332].
Organelles have been found to be enriched in specific species of PIP's – e.g. the plasma
membrane is highly enriched in PI(4,5)P2, whereas MVB's and early endosomes are enriched
in PI(3)P, and the Golgi is highly enriched in PI(4)P [331, 333].
PIP levels at organelles are maintained by a variety of phopsphatidylinositol kinases (PIK's)
and phosphatases. Each mediates the phosphorylation/dephosphorylation of a defined
position on the inositol head group [236, 316, 334‐336]. PIK's and phosphatases maintain
PIP's that are not always located on the same membrane as the enzyme, e.g. a substantial
part of the PI(4)P PM pool is supplied by type III PI‐4 Kinase (PI4KinIIIα) that resides in the

ER, where exchange may occur at various contact sites between the ER and the PM [337‐
339]. A recent report has also identified a PI4KinIIIα population that is present at the PM
which also influences the PM localized amount of PI(4)P [340]. Similarly, PI(4)P
dephosphorylation at the PM is mediated by the ER resident Sac1 phosphatase at ER‐PM
contact sites [339, 341‐344].
PI modifiers are regularly targeted to specific subdomains by small GTPases, e.g. Rac1
45
recruits the lipid phosphatase synaptojanin 2 to the PM [345]. Small GTPases also function
as activators of the PI modifiers. For instance, Rab5 activates the type III phosphatidyl‐3‐
kinase at endosomes [346].
Several protein motifs have been identified that recognize and bind PIP's. These include
FYVE, PX, PH, ENTH, ANTH, Tubby, FERM and DDHD domains [347‐357]. These motifs are
found in numerous proteins with varied functions such as cytoskeletal remodeling, protein
sorting, vesicular trafficking, and lipid metabolism [347‐357].
Golgi and PI(4)P
PIP signaling within the early secretory pathway and Golgi is mainly associated with PI(4)P
[162, 358, 359]. PI(4)P is highly enriched in the Golgi, where the lipid is integral in transport
signaling and transport initiation from all compartments of the organelle [331].
The Golgi pool of PI(4)P is supplied either by lipid transfer protein shuttling the from ER to
Golgi, e.g. by PITPβ [360], or by kinase activity through the two kinases PI4KinIIα, and Arf 1
associated PI4KinIIIβ [338, 361].
PI(4)P also recruits a family of Four‐Phosphate‐Adaptor Proteins (FAPP)‐Related proteins
FAPP1 and FAPP2 through their PH domains [362, 363]. The FAPP proteins promote
transport from Golgi to the PM by supporting Golgi vesicle formation [364].
The presence of PI(4)P at the cis‐Golgi is furthermore necessary for the assembly of the
trans‐SNARE complex and the subsequent fusion of COPII vesicles with Golgi acceptor
membranes [359].
PI(4)P and ERES formation
The formation of ERES has proven to be dependent upon local increases of PI(4)P
concentrations [162, 184, 358]. The majority of PI(4)P synthesized in the ER is formed by

46
phosphorylation of PI through two PI‐4 Kinases, the ER membrane abundant PI4KinIIIα and
PI‐4 Kinase II α (PI4KinIIα) that associates with membranes throughout the cell [236, 338,
361, 365, 366].
Initial experiments using yeast COPII components showed a dependence of lipid‐
composition for the nucleotide‐induced recruitment of the COPII proteins to proteolipsome
[184]. It was found that Sar1p could bind to liposomes composed of 53 mol% PC, 23 mol%
PE, 8 mol% PS, 5 mol% PA and 11 mol% PI at nearly normal levels, but recruitment of
Sec23/24p and Sec13/31p was weak. Replacing a portion of the PI with PI(4)P dramatically
increased the Sar1p‐dependent recruitment of Sec23/24 and Sec13/31. Inclusion of
PI(4,5)P2 and CDP‐DAG further enhanced the binding of the coat proteins. Recruited Sar1p
levels did not increase significantly in response to the changes in lipid composition,
suggesting that the PIP's are involved in mediating and stabilizing the recruitment of the
coat layers [184].
These observations have been supported by later experiments, where a decrease in Sar1‐
dependent recruitment of Sec23 was observed in budding assays performed in conditions
with low lipid kinase activation due to a reduction in the available pool of ATP [162]. The
Sar1 activated recruitment of Sec23 was shown to be ATP dependent, but could be
rendered ATP independent by supplying the reaction with PI(4)P micelles. Similar effects
were also observed when localizing COPII component in morphological transport assays
using semi‐intact cells. Addition of GST‐Fapp1‐PH to the reaction markedly reduced Sar1‐
induced nucleation of both the Sec23/24 layer as well as the Sec13/31 layer. These results
show that COPII nucleation and assembly is dependent on the presence of PI(4)P [162].
Depletion of the PI(4)P pool at the ER, by knock‐down experiments targeting PI4KinIIIα, has
been shown to decrease the number of visible ERES in HeLa cells [358]. A reduction in the
transport efficiency of VSV‐G could also be measured. When PI4KinIIIα was knocked down in
Brefeldin A (BFA) treated cells, a reduction in spot intensity and size of GFP‐tsO45‐VSV‐G‐
marked ERES was observed [358, 367]. The authors went on to mimic chronic increase in
cargo load, by overexpression of the anterograde GABA transporter 1 (GAT1). This
treatment caused an app. 30 % increase in the number of visible ERES. siRNA‐mediated
reduction of PI4KinIIIα did not influence the relative increase in ERES numbers, ruling out an
influence of PI4KinIIIα on de novo ERES formation during chronic cargo load. The factors

promoting PI4KinIIIα‐mediated elevation of PI(4)P at ERES have yet not been identified
[358].
47
Local PI(4)P increases have been observed at ERES during loading of VSV‐G‐tsO45 [162]. In
vitro budding assays from microsomes isolated from cells overexpressing ts‐O45‐VSV‐G
showed a clear inhibition of ER export when GST‐Fapp1‐PH was added to the reactions [162,
368]. Adding PH domains targeting PI(3,4,5)P3 did not modulate the ts‐O45‐VSV‐G export,
whereas only high concentrations of PI(4,5)P2‐binding PH‐domains showed an inhibitory
effect. Furthermore, addition of liposomes composed with PI(4)P (but not PI‐free liposomes)
together with the GST‐Fapp1‐PH fragment to the budding reaction maintained the normal
levels of ER‐export, implying that GST‐Fapp1‐PH inhibited the ER export due to its PI(4)P
binding capabilities [162]. The inhibitory effect of GST‐Fapp1‐PH was repeated when added
to semi‐intact cells during induced morphological GFP‐tsO45‐VSV‐G transport assays [162,
319]. The specificity of PI(4)P at ERES was verified by addition of the cytosolic domain of the
PI(4)P preferring yeast phosphatase Sac1 to both the budding reaction and the
morphological transport assay. In both instances, ER export was inhibited [162, 369].
The levels of PI(4)P could be shown to increase in a cyclic manner at 3 min and 9 min in
response to Sar1‐GTP activation of ERES formation on purified rat liver microsomes. Similar
transient activity had been observed previously with Arf1 on Golgi membranes, and was
presumed to reflect limiting PI levels that are dynamically regenerated at the membranes
[162, 370]. Sar1‐GTP activation in the presence of the phosphatase inhibitor orthovanadate
led to a 4‐fold increase of PI(4)P and PI(4,5)P2 levels. This implies that Sar1 does not control
PIP levels at ERES by inhibition of phosphatase activity. Rather, Sar1 stimulates kinase
activity at the ERES. This furthermore implies that Sar1 provides a coupling between COPII
assembly and PI(4)P formation [162].
Sar1‐GTP induces ERES tubulation in semi‐intact cells under cytosol free reaction conditions,
in which cargo such as ts‐O45‐VSV‐G is selectively concentrated [157]. PI(4)P could be
visualized as present in these structures. The Sar1‐GTP induced tubulation could
furthermore be inhibited by addition of GST‐Fapp1‐PH. This implied that PI(4)P may also
support Sar1‐induced tubule constriction at ERES [162].
Why ER budding needs locally elevated PI(4)P levels still has to be fully investigated. The

48
recruitment of yeast COPII components to synthetic liposomes suggests a role in assembling
and stabilizing the formation of the COPII coat [184]. These interactions are though
redundant in the presence of Sec16p. Studies in yeast furthermore show that depleting the
major PI4‐kinases does not impede ERES budding, but rather inhibits the fusion of COPII
vesicles with Golgi acceptor membranes [359, 371]. In the mammalian system COPII vesicles
fuse homotypically close to the bud site and prior to merging with the Golgi [50, 51]. Taken
together, these results imply that PI(4)P play a more important role for maintaining the
COPII linkage and possibly the subsequent recruitment of SNARE proteins to the vesicle.
Indeed, in vitro experiments where COPII assembly at ERES was induced by introduction of
Sar1 seem to imply that the lipid may act more as a signal for recruitment of accessory
proteins that can maintain the scaffolding organization during budding [159, 162, 358].
The formed lipids and in particular PI(4)P need to be decoded at the ERES. Here, two
proteins have been identified with a potential role in responding to the lipid signals, Sec16
and p125A.
Sec16
Maintenance of the transitional domain at ERES has been associated with the Sec16
proteins. Sec16 was early on found to bind to Sec23, 24 and 31, implying a role in the
assembly of the COPII vesicle [136‐138, 372‐375]. A point mutation in the yeast Sec16
(Sec16p), called dot1, was observed to cause breakdown and dispersion of the otherwise
unique and defined tER in Pichia pastoris (P. pastoris). This organism stands out compared
to Saccharomyces cerevisiae (S. cerevisiae) by actually having stacked Golgi cisternae ‐
almost resembling a mammalian Golgi ‐ in juxtaposition to specified tER/ERES. The
breakdown furthermore caused a dispersion of the coalesced Golgi, indicating that the P.
pastoris Sec16 plays a role in the maintenance of the cohesion between Golgi stacks. This
could be effected either by tethering individual compartments to each other, or through the
innate role of Sec16 in controlling the integrity of the tER and influence the transport
dynamics needed to maintain a collected Golgi [376].

Sec16 structure
49
Two mammalian homologs of Sec16 (Sec 16 A and B) have been identified. Sec16A is a 250
kDa protein that most closely resembles the S. cerevisiae ortholog functionally. Sec16A
contains a central conserved domain (CCD) and a C‐terminal region of app. 250 residues that
is conserved amongst orthologs [376‐378]. Sec16B is a shorter and less characterized 117
kDa protein, which contains a CCD but, compared to Sec16A, has a substantial truncation at
the N‐terminus and lacks the conserved C‐terminal region [378‐381] (see fig. 11).
Figure 11 – Graphical overview of Sec16A and B protein structure – Top: Sec16A, a protein of 2110 residues. Sec16A
contains a Central Conserved Domain (CCD), a highly charged arginine rich sequence (RRS), and finally a conserved C‐
terminal domain of app. 250 residues. Bottom: Sec16B, a protein of 1061 residues, which is a shorter homolog of Sec16A
that contains a CCD. Whether Sec16B utilizes an RRS similarly to Sec16A needs to be further investigated.
Depletion of either Sec16A or Sec16B has been shown to inhibit ER‐to‐Golgi transport [377,
378], but only Sec16A has been identified to directly associate with COPII components ‐
Sec23 and Sec13 ‐ through the conserved C‐terminal domain [378, 382, 383].
Both human Sec16 proteins have been observed to localize with tER/ERES. Work using the
Drosophila melanogaster (D. melanogaster) ortholog ‐ dSec16 ‐ identified an arginine‐rich
region at the N‐terminus that, in conjunction to the Conserved Central Domain, was needed
to localize the domain to the tER [384]. In agreement, experiments using N‐terminal
deletions in mammalian Sec16A caused an apparent loss of tER targeting [378]. Later
observations with a different, longer, version of the mammalian Sec16A, mapped the tER
targeting to the CCD and a highly charged region enriched in arginine (RRS) as observed for
dSec16 [383].
The mammalian Sec16A and Sec16B have both been reported to oligomerize [378]. Recent
reports identified sequence similarities within the CCD to the ACE1 in Sec31, and, in
agreement, Sec16A has been crystallized in complex with Sec13 in a similar β‐propeller‐
ACE1 interaction to that observed between Sec13‐Sec31A (see fig. 6) and Sec13‐NUP145C

50
[385]. The crystal structure furthermore revealed that Sec16A and Sec13 form a tetrameric
complex (see fig. 12). The inner core of the complex consists of the trunks of the two Sec16s
forming a curved solenoid backbone. Sec13 associates at each end of the Sec16A dimer core
[385]. It should also be noted that the initially mentioned dot 1 mutation was mapped to the
ACE1/CCD.
The implications of these findings are still not fully understood, but the authors
hypothesized that a Sec13/Sec16A complex could act as a membrane tethered scaffold that
initiates the recruitment and stabilization of the inner layer of the COPII coat (Sec23/24).
The Sec13/16A scaffold would subsequently get displaced by the Sec13/31A cage [376, 385].
Sec16 functions
A number of the observations done on Sec16 provide evidence for the protein's
involvement in both the nucleation of novel ERES and maintaining existing ERES. This
conclusion is based on examining Sec16 from several eukaryote systems ranging from yeast
to humans.
Figure 12 – Sec16A/Sec13 Tetramer
Complex ‐ Sec16A (green and light blue)
in complex with Sec13 (yellow and blue)
seen from two angles. Folding of the
Sec16A C‐terminus ACE1 associated
"trunks", forms a curved solenoid
backbone (green and light blue). Sec13
(yellow and dark blue) forms separate
compact ball structures in extension to
the Sec16A solenoid backbone. Modeled
from PDB accession # 3MZK‐ Sec13‐Sec16
Tetramer – using 3D‐molecule viewer
(Invitrogen).

51
Evidence for Sec16 acting as an initial scaffold for the COPII nucleation has been observed
both in vitro and in vivo. S. cerevisiae Sec16p has been shown to bind to synthetic liposomes
in response to Sar1 activation, and aid in the recruitment of the outer and inner layers of
the COPII coat. Sec16p has furthermore been observed to decelerate the rate of Sec31
increased GTP hydrolysis in Sar1 when associating with Sec23 on microsomes [375]. Sec16p
has also been recognized to stabilize the cage complex post‐GTP hydrolysis on synthetic
liposomes, and thereby inhibit premature disassembly. These observations imply that
Sec16p acts as a scaffold during vesicle coat assembly [185]. The same study furthermore
showed that Sec16p targeted and bound liposomes with acidic lipid composition
independent of Sar1p activation. Substituting the acidic lipid composed liposomes with non‐
acidic lipid composed liposomes inhibited the Sar1p‐independent binding of Sec16p.
Instead, Sec16p was recruited in response to Sar1p activation, and this in turn led to
recruitment and stabilization of the inner and outer layer. Sar1p activation on non‐acidic
liposomes in the absence of Sec16p did not lead to the recruitment of Sec23/24 and
Sec13/31. This suggests that Sec16p can overcome the necessity of acidic lipids for ERES
nucleation, and act as a scaffold for the stabilization of the COPII coat [185].
In vivo results in mammalian systems have observed that Sec16A remains associated with
tER during mitosis in contrast to the COPII coatomers that become largely cytosolic [386].
Sec16A wt was observed to nucleate COPII at tER during exit from metaphase, whereas
deletion mutations inhibiting Sec16A association with Sec23A caused dispersed recruitment
of Sec23A. These findings indicate that the mammalian Sec16A , equivalent to Sec16p, also
plays a role in defining and initiating the assembly of ERES [386].
Investigations of the dynamics of Sec16A at ERES show the protein to have role in
maintaining and controlling the ERES during ER export. This function has been observed
both at steady state transport levels as well as during conditions of cellular stress due to
elevated levels of protein expression and imposed export pressure.
FRAP studies of the protein showed that Sec16A has a generally slower recycling time
compared to Sec23 [383]. The same study used EM tomography to localize the binding of
Sec16A to the outer edge of distinct cup‐like structures on the ER membrane. Interestingly,
the tomography showed clear spatial separation between Sec16A and Sec31A. It was further

shown that blocked recycling of Sec23/24 still maintained Sec16A localizing to distinct
52
puncta, and did not alter the rate of Sec16A recycling [383].
The function of Sec16A has also been shown to play a pivotal role in responding to both
acute and chronic increases in cargo load [358]. In the acute situation, cells normally
respond by coalescing ERES into fewer and larger puncta, probably due to elevated local
COPII assembly in response to the increased protein production. Knock‐down of Sec16A
decreased the number of ERES, and when acute cargo load was induced, using BFA and
followed by washout, no increase in ERES numbers nor size was observed. This indicated
that Sec16A is a necessary component in response to acute increases in cargo load [358,
367]. During chronic elevated cargo load, the number of observed ERES was increased and a
marginal increase in Sec16A expression was also recorded. In an effort to get a better
Sec16A response, levels of the protein were lowered with siRNA before a chronic cargo load
was introduced by overexpression of a cargo protein. These results showed a clearly
induced elevation of Sec16A in response to the chronic load. The higher levels of Sec16A
could be linked to the induction of the Unfolded Protein Response. This implies that a
Sec16A threshold has to be reached, and that this threshold usually is present during regular
steady state levels of the protein for the handling of chronic cargo loading [358].
Various roles for Sec16, which might not necessarily translate to the mammalian system,
have been found in orthologous systems. As described in the following, these tasks indicate
that Sec16 functions are controlled and regulated in response to changes in the local cellular
environment.
A kinase‐depletion screen in the D. melanogaster identified dSec16 to be regulated by the
Extracellularly Regulated Kinase 7 (ERK7) in response to serum or amino acid starvation
[387]. The amino acid depravation caused a stabilization of ERK7 as well as disassembly of
the tER. Overexpression of ERK7 was furthermore shown to cause dispersal of dSec16.
These results imply a tight regulation between dSec16 and the establishment of highly
productive tERs [387].
Sec16 in Caenorhabditis elegans (C. elegans) (Sec16(Ce)) was recently shown to interact
with Tyrosine Receptor Kinase Fused Gene‐ 1 (TFG‐1), a factor required for protein secretion
in the nematode [388]. TFG‐1 was shown to aid Sec16(Ce) accumulation at ERES. The TFG‐1

hexamer was further shown to influence Sec16(Ce) complex assembly. It should be noted
that Sec16(Ce) did still form ERES‐like structures in TFG‐1‐depleted cells, whereas TFG‐1
seemed to aggregate in Sec16‐depleted cells. A following co‐immunoprecipitation
experiment from transient expressions in HeLa cells with the N‐terminus of the human
53
homolog of TFG‐1 and an mCherry‐tagged human Sec16B, showed that these two proteins
do interact, adding another possible regulator in the ERES maintenance [388].
A C‐terminal fragment of Sec16p (565‐1235) is capable of delaying the Sec31‐induced GTP
hydrolysis in Sar1 [176]. The authors hypothesized that coupling of Sec16p with the
coatomer, likely with Sec24, would ensure that only mature vesicles with loaded cargo
would be released from the bud site [176]. It should also be noted that the C‐terminus of
Sec16p has been identified to bind the Sec12p homolog Sed4p, which is known to stimulate
Sar1p‐GTP hydrolysis in response to absence of cargo [165, 166, 389]. The Sec16p influence
on GAP activity was further substantiated by a recent report showing that a fragment of
Sec16p (1639‐2195) can compete with full lengthSec16p during budding. The fragment was
shown to cause a substantial delay in the Sec23 mediated GAP activity [375]. In vitro
budding assays on artificial liposomes demonstrated that the Sec16p (1639‐2195) inhibited
the recruitment of Sec31 to the ERES and thus inhibiting the Sec31 mediated catalysis of the
Sar1‐GTP hydrolysis. These add to the function of Sec16p a role as a monitor of vesicle
maturation.
Sec16B
The functions of Sec16B have still not been fully investigated. Sec16B seems to exist in a
larger complex together with Sec16A, and targets towards ERES through an N‐terminal
region [378].The protein was originally identified as potentially binding to the regucalcin
promoter [379]. It was later suggested that Sec16B enhances regucalcin expression in
response to cytokines and various Ca 2+ signaling factors [379, 390, 391]. How exactly Sec16B
regulates regucalcin has still not been fully determined [379, 390, 391]. Depletion of Sec16B
has a disruptive effect on tER morphology and inhibits export of a GFP‐tagged Golgi protein,
GalNac‐T2‐GFP, from the ER. However, as mentioned above, Sec16B does not comprise the
C‐terminal region conserved among Sec16A orthologs involved in binding to Sec23 and
Sec13, and has not been identified to associate with any COPII components [378].

Still, Sec16B has been identified as important in the budding of specialized vesicle
compartments from the ER in the mammalian system. Sec16B was recently shown to
54
participate in the peroxisome biogenesis [392]. Depletion of Sec16B, but not Sec16A, caused
changes in peroxisome morphology and changed the distribution of peroxisomal membrane
proteins. These changes in distribution could be attributed to a transport inhibition of the
peroxisome‐specific transport receptor Pex16, implying a specific role for Sec16B in a
peroxisome‐dedicated ER export machinery [392].
Overall, the mechanistic roles of the Sec16 proteins are presently not well understood.
Apparent functional differences between the protein in yeast and higher eukaryotes seem
to indicate that roles maintained by a single protein in yeast and lower order organisms may
have been split up in later evolutionary stages to support more elaborate physiological
demands for cargo regulation, unique structural ERES morphology and overall ERES
organization in higher eukaryotes.
p125A (Sec23IP)
The mechanisms that regulate ERES assembly especially in response to signals in the local
lipid environment are still unknown. In this thesis we hypothesize that p125A – also known
as Sec23IP – might be one plausible regulator of ERES that utilizes selective lipid recognition.
p125A architecture
p125A – a 125 kDa (1000 aa) protein (see fig. 13) – was identified in GST pull‐down assays
using GST‐Sec23 [354]. p125A consists of an N‐terminal WWE motif, a central non‐functional
lipase motif, followed by a domain with a sterile α‐motif (SAM domain) and a DDHD (Asp‐
Asp‐His‐Asp) domain at the C‐terminus [393]. Additionally, p125A comprises a proline‐
glutamine (P‐Q) rich N‐terminal domain, to which Sec23 binding has been localized. More
specifically, Sec23 binding was mapped to residues 135‐259 [393, 394].

WWE motifs are named after the characteristic two conserved tryptophans and a single
glutamic acid residue [395]. They are also found in several E3 ubiquitin ligases, poly‐ADP‐
55
Figure 13 – Graphical overview of p125A – The 1000 residues long p125A consists of an N‐terminal proline – glutamine
(P‐Q) rich region (yellow) with a WWE motif at the end (WWE). The P‐Q region has been identified to associate with
both Sec23 and Sec31. It contains a non‐functional lipase motif (black) in front of a putative oligomerizing SAM domain
(blue) in the central part of the protein. Finally, p125A has a DDHD domain (Bordeaux red) at the C‐terminus that is
presumed to provide the proteins lipid recognition and binding activity.
ribose polymerases and in p125B [395, 396]. There is a low degree of sequence homology
within this domain family, which at the structural level superficially resembles ubiquitin. The
WWE motifs are presumed to mediate protein‐protein interactions that promote poly‐ADP‐
ribosylation or ubiqutination [395, 396].
SAM domains are among the most abundant protein‐protein interaction motifs known [397,
398]. They are mostly found in the context of larger multidomain proteins located in all
cellular compartments mirroring the participation in a wide variety of processes [398]. A
general commonality is their capability to modulate function by homo‐ or hetero‐
oligomerization [399‐402].
p125A is a homolog of DDHD1 (previously known as phosphatidic acid preferring‐
Phospholipase A1 (PA‐PLA1)) [354, 355], and belongs to a family of proteins defined by a
conserved DDHD motif. The family consists of p125A, DDHD1 and a smaller homolog of
p125A called p125B.
p125B
p125B consists of a C‐terminal DDHD domain, an N‐terminal WWE motif, and a centrally
located lipase motif followed by a SAM domain (see fig. 14).
Figure 14 – Graphical overview of p125B ‐ The 711 residues long p125B contains an N‐terminal WWE motif (red) whose
binding target has not been determined. p125B furthermore comprises of a central lipase domain (black) with activity
that targets several phospholipids, followed by a SAM domain (blue) that has been identified to cooperate with the C‐
terminal DDHD domain (Bordeaux red) in targeting and binding to lipids. The DDHD domain functionality has been
identified as essential for p125B's lipase activity.

56
In contrast to p125A, p125B does not bind to Sec23A [393, 403, 404]. Although p125B has
several domains in common with p125A, and appears to target membranes enriched in
specific lipids, the protein does not seem to have the same functions as p125A.
p125B has been shown to target and bind at the cis‐Golgi [405]. The protein has also been
shown to have PLA1 activity with a preference towards hydrolyzing PA. Minor activity is
observed against PS and PC in the presence of Triton X‐100, and towards PE in detergent
free conditions [403]. The lipase activity has been shown influential for the targeting
towards the Golgi [405]. The localization has been further shown to be influenced by Sac1
phosphatase activity [393].
The mechanism for p125B targeting appears to also influence the protein's lipase activity, so
how does p125B recognize specific lipids? A recent study has identified the DDHD domain of
p125B as part of a monophosphorylated PI binding element together with the SAM domain
[393]. The study showed that purified full length GST‐tagged p125B recognized PI(3)P, PI(4)P
and PI(5)P in a lipid blot over‐lay assay. Furthermore, a marked increase in targeting
towards PI(4)P and PI(4,5)P2‐containing liposomes was detected using a lipase‐inactive
mutant of the purified protein. As potential lipid binding capabilities by SAM domains have
also been reported [406, 407], and since a SAM domain is present in both p125A and p125B,
the authors reasoned that the present SAM domain might either confer PIP binding, or
collaborate with the downstream DDHD domain for targeting and lipid binding [393].
Purified GST‐tagged p125B (SAM) domain, p125B (DDHD) domain and a C‐terminal fragment
comprising both the p125B (SAM) and the p125B (DDHD) domain – p125B (SAM‐DDHD) –
were examined by lipid blot over‐lay [393]. It was found that neither SAM alone nor DDHD
alone conferred any lipid targeting. However, the SAM‐DDHD containing fragment showed
targeting towards PI(4)P, implying that this module was responsible for the p125B targeting
towards the Golgi. Alanine substitutions of a positively charged arginine, lysine and alanine
(RKA) group in the SAM domain caused the full‐length protein to lose Golgi targeting,
supporting a role for the SAM domain in membrane targeting of p125B. In addition, the
DDHD domain was shown to be essential for the p125B lipase activity, as alanine
substitutions of the actual DDHD motif abolished the PLA1 activity markedly [393]. It should
be noted that this group did not analyze the structural effects of the RKA mutation and as
consequence the group did not define a specific role for the SAM domain [393].

A more recent genetic study has recognized several mutations in p125B that causes a
recessive form of complex spastic paraplegia as well as intellectual disabilities [408]. The
majority of the defects cause frame shifts truncating or abolishing the DDHD region. One
57
interesting point mutation influences a conserved RIDYXL motif found throughout the DDHD
family of proteins causing the arginine to be substituted with a histidine. Cerebral proton
MRS of affected individuals showed an abnormal lipid peak similar to a characteristic lipid
peak found in individuals afflicted with Sjögren‐Larssen syndrome, indicative of abnormal
brain lipid accumulation [408, 409]. These observations imply that p125B plays an important
role in the normal development of the central nervous system (CNS) [408].
Cellular Localization of p125A
p125A is expressed ubiquitously in an expression pattern similar to Sec23 [354]. p125A's
homology to p125B, and the presence of both a lipase motif and of a DDHD domain that is
known to bind lipids, implies that p125A may also possess similar specific lipid recognition.
But, as will become apparent, p125A and p125B do not seem to target the same type of
membranes.
Transient expression of p125A showed that it co‐localized with ERGIC53 and β‐COP to VTC's
[354]. Further dissection, using a mAb raised against the first 134 residues of the protein,
has specified a strong perinuclear co‐localization with the COPII components Sec31 and
Sec23, and a lesser overlap with β‐COP, clearly indicating a role at ERES. EM analysis showed
that p125A does not localize within Golgi stacks, yet resides in regions between the ER and
Golgi in a pattern similar to Sec31 [354].
Inhibition of retrograde transport by BFA treatment does not disperse p125A in a similar
manner as the rapidly recycling ERGIC53. Instead, the protein retains its perinuclear co‐
localization with Sec31. Expression of Sar1 (H79G) furthermore clusters Sec23, Sec31 and
p125A at perinuclear ERES. Additionally, recruitment of p125A to purified ER microsomes is
Sar1A dependent, indicating that p125A is recruited as part of the COPII complex [410].
Consequences of modulating p125A expression levels
So what type of influence does p125A confer at the ERES? During overexpression of p125A,
ERES perturb and the overexpression induces disorganization of both VTC's and the cis‐
Golgi, which causes the VTC's and subsequently the Golgi to disperse [354, 394, 410]. Under

these conditions, p125A collects into larger perinuclear structures that contain ERES
components such as Sec23, Sec31, p115 and GM130, whereas β‐COP and ERGIC53 are
58
dispersed. This has been interpreted as p125A being capable of inducing organized cellular
localization of the ERES with the ERGIC and the cis‐Golgi as a result of ectopical expression.
p125A also influences the later compartments of the Golgi, e.g. medial‐, trans‐Golgi and the
TGN. siRNA‐induced p125A depletion disperses the Golgi, yet it maintains its cis‐trans
organization. This implies that p125A knock‐down compromises the stacking and fusion
necessary for Golgi ribbon formation. Knock‐down also causes dispersion of the usually
perinuclear‐concentrated and ERES‐associated Sec23 and Sec31 within 48 hours. After an
additional 24 hours, β‐COP becomes dispersed, indicating that p125A plays a role in
maintaining the cellular distribution of ERES at an early stage of ERES formation [410, 411].
The p125A association with the COPII components and its influence on the organization of
the compartments within the biosynthetic transport pathway means that ER export is also
affected. Traffic delays are observed in p125A‐depleted cells when monitoring VSV‐G‐tsO45‐
GFP transport. Transport inhibition was also observed for secretion of alkaline phosphatase.
Both findings indicate an accessory role for p125A in influencing COPII‐mediated ER export.
Since transport is still ongoing albeit delayed in knock‐down cells, implications are that
p125A is an essential ERES regulator involved in improving the efficiency of COPII‐mediated
cargo export from the ER [411]. These effects are almost identical to the effects observed
during knock‐down of Sec16A, where steady state trafficking is also perturbed [358].
These observations have been translated into biological systems. p125A knock‐out mice are
viable, but show impaired male fertility due to defective spermiogenesis. Specifically,
spermatids lacked acrosomes, an organelle covering part of the head of the sperm
containing the enzymes responsible for dissolving the zona pelucida of the ovum. The
formation of the acrosome is mediated by fusion of pro‐acrosome vesicles derived from the
trans‐Golgi. The specific role of p125A in spermiogenesis has yet to be determined [412].
p125A ERES targeting and interactions
Targeting of p125A to ERES is dependent on both the protein's association with ERES
protein components as well as its targeting to specific lipids. Targeting of p125A to lipids has
prior to the paper of this thesis not been fully resolved.

59
Preceding observations with a fragment comprising the N‐terminal 259 residues of p125A,
containing the P‐Q rich region and the Sec23 binding domain, showed targeting to ERES.
This is also the case when the phospholipase motif is expressed without the P‐Q‐domain
[394]. Chimeric substitution of the DDHD domain with the DDHD of p125B still retained
targeting towards ERES, whereas substituting the DDHD domain of p125A with the DDHD
domain from DDHD1 did not retain targeting, but rather dispersed the Sec23 expression
pattern. These results imply that specific phospholipid recognition is necessary for ERES
localization of p125A [403, 405, 410]. Importantly, p125A appears to lack lipase activity
[382, 393, 403]. However, a recent study has shown that a purified GST‐tagged full‐length
p125A targets and binds PI(3)P, PI(4)P and PI(5)P in lipid blot over‐lays [393].
Additional parts of p125A are also involved in associating with COPII components. Pull‐
downs of p125A with a GST‐tagged fragment of Sec31A (1041‐1220) show that p125A
interacts directly with Sec31A [411]. In gel filtration assays, p125A co‐elutes with Sec31A
and Sec13 but not with Sec23, implying that Sec13/Sec31/p125A co‐exist as a complex in
the cytosol. This finding may also explain the unexpectedly high molecular weight that has
been seen in previous reports of the Sec13/31 complex in gel‐filtration assays. The
theoretical MW of the complex is app. 370 kDa, but the components elute as a 600‐700 kDa
complex [169, 411, 413‐415]. Immunodepletion of p125A caused a proportional depletion
of Sec31A, indicating that the two proteins are likely bound together in the cytosol. The
region responsible for binding to Sec31A has been mapped to residues 260‐600 of p125A
and comprises the end of the P‐Q domain containing the WWE motif (259‐342). The Sec31A‐
p125A association was also inferred by live‐cell imaging experiments, where the two
proteins co‐localized in dynamic vesicular structures capable of undergoing homotypic
fusion for time periods of more than 30 min [411].
Taken together, these studies indicate that p125A is recruited to the ERES together with the
outer COPII coat layer, likely as an integral part of the Sec13/31 complex. At the ERES,
p125A binds Sec23 and thus promotes the stabilization and scaffolding of the COPII coat.
COPII coat stabilization is further promoted by the binding of p125A to a specific sub‐set of
phospholipids.

p125A and disease
Since p125A appears to be an important part of the COPII complex, various pathological
diseases may be associated with p125A dysfunction. Indeed, a few diseases have been
linked to variations in p125A expression levels.
p125A has been identified as a potential candidate involved in Waardenburg Disease that
60
causes craniofacial dysmorphy due to defects in the developing neural crest [416]. The study
identified p125A through screening and comparing orthologous disease phentoypes
between negative gravitropism (growth direction) defects of Arabidopsis Thaliana
translated to vertebrate systems. Indeed, Xenopus p125A is prominently expressed in
migrating neural crest cells in embryos. Morpholino experiments targeting p125A
unilaterally caused marked defects in the neural crest migration pattern at the injection
side, corroborating the results of the screen [416].
The p125A gene has a chromosomal position at a region that has quantitative trait loci (QTL)
for femur strength in a specific cross of inbred rats. Thus, low femur strength showed a
strong correlation with lower levels of p125A expression in these breeds [417].
Both the observed morpholino p125A migration defect and the influence of p125A on rat
femur strength imply that p125A influences collagen secretion. These observations are in
agreement with studies showing that defects in COPII affect collagen secretion [213, 214,
418].
Knock‐down of the p125A and p125B ortholog CG8552 in D. melanogaster did not cause
visual phenotypical abnormalities in the flies [408]. This study did though observe a mild but
highly significant decrease in chemical synapses, a.k.a. active zones, in the observed
individuals. These observations indicate the p125A and p125B might play an important role
in the development of the CNS.
Recently, p125A gene expression was identified as a possible effector in melanoma
progression [419]. Superficial Spreading Melanoma (SSM) showed a lower degree of p125A
mRNA expression compared to Nodular Melanoma (NM), which correlated with frequent
deletions of the p125A gene. NM has a higher rate of re‐occurrence and does not show a
high degree of initial downward‐stage migration compared to SSM, but the overall role of
p125A in carcinogenesis needs to be defined [419].

61
References
1. Golgi, C., Intorno Alla Struttura Delle Cellule Nervose. Bolletino Della Società Medico‐
Chirurgica Di Pavia, 1898(13): p. 3‐16.
2. Jamieson, J.D. and G.E. Palade, Intracellular transport of secretory proteins in the pancreatic
exocrine cell. I. Role of the peripheral elements of the Golgi complex. J Cell Biol, 1967. 34(2):
p. 577‐96.
3. Neutra, M. and C.P. Leblond, Synthesis of the carbohydrate of mucus in the golgi complex as
shown by electron microscope radioautography of goblet cells from rats injected with
glucose‐H3. J Cell Biol, 1966. 30(1): p. 119‐36.
4. Neutra, M. and C.P. Leblond, Radioautographic comparison of the uptake of galactose‐H and
glucose‐H3 in the golgi region of various cells secreting glycoproteins or
mucopolysaccharides. J Cell Biol, 1966. 30(1): p. 137‐50.
5. Porter, K.R., A. Claude, and E.F. Fullam, A Study of Tissue Culture Cells by Electron Microscopy
: Methods and Preliminary Observations. The Journal of experimental medicine, 1945. 81(3):
p. 233‐46.
6. Porter, K.R., Observations on a submicroscopic basophilic component of cytoplasm. J Exp
Med, 1953. 97(5): p. 727‐50.
7. Siekevitz, P. and G.E. Palade, A cytochemical study on the pancreas of the guinea pig. I.
Isolation and enzymatic activities of cell fractions. J Biophys Biochem Cytol, 1958. 4(2): p.
203‐18.
8. Siekevitz, P. and G.E. Palade, A cytochemical study on the pancreas of the guinea pig. II.
Functional variations in the enzymatic activity of microsomes. J Biophys Biochem Cytol, 1958.
4(3): p. 309‐18.
9. Siekevitz, P. and G.E. Palade, A cyto‐chemical study on the pancreas of the guinea pig. III. In
vivo incorporation of leucine‐1‐C14 into the proteins of cell fractions. J Biophys Biochem
Cytol, 1958. 4(5): p. 557‐66.
10. Siekevitz, P. and G.E. Palade, A cytochemical study on the pancreas of the guinea pig. 5. In
vivo incorporation of leucine‐1‐C14 into the chymotrypsinogen of various cell fractions. J
Biophys Biochem Cytol, 1960. 7: p. 619‐30.
11. Caro, L.G. and G.E. Palade, Protein Synthesis, Storage, and Discharge in the Pancreatic
Exocrine Cell. An Autoradiographic Study. J Cell Biol, 1964. 20: p. 473‐95.
12. Voeltz, G.K., M.M. Rolls, and T.A. Rapoport, Structural organization of the endoplasmic
reticulum. EMBO reports, 2002. 3(10): p. 944‐50.
13. Vedrenne, C., D.R. Klopfenstein, and H.P. Hauri, Phosphorylation controls CLIMP‐63‐
mediated anchoring of the endoplasmic reticulum to microtubules. Mol Biol Cell, 2005. 16(4):
p. 1928‐37.
14. Grigoriev, I., et al., STIM1 is a MT‐plus‐end‐tracking protein involved in remodeling of the ER.
Curr Biol, 2008. 18(3): p. 177‐82.
15. Friedman, J.R., et al., ER sliding dynamics and ER‐mitochondrial contacts occur on acetylated
microtubules. The Journal of cell biology, 2010. 190(3): p. 363‐75.
16. Terasaki, M., L.B. Chen, and K. Fujiwara, Microtubules and the endoplasmic reticulum are
highly interdependent structures. J Cell Biol, 1986. 103(4): p. 1557‐68.
17. Voeltz, G.K., et al., A class of membrane proteins shaping the tubular endoplasmic reticulum.
Cell, 2006. 124(3): p. 573‐86.
18. Hu, J., et al., Membrane proteins of the endoplasmic reticulum induce high‐curvature tubules.
Science, 2008. 319(5867): p. 1247‐50.
19. Shibata, Y., et al., The reticulon and DP1/Yop1p proteins form immobile oligomers in the
tubular endoplasmic reticulum. The Journal of biological chemistry, 2008. 283(27): p. 18892‐
904.

62
20. Shibata, Y., et al., Mechanisms determining the morphology of the peripheral ER. Cell, 2010.
143(5): p. 774‐88.
21. Becker, F., et al., Expression of the 180‐kD ribosome receptor induces membrane
proliferation and increased secretory activity in yeast. The Journal of cell biology, 1999.
146(2): p. 273‐84.
22. Puhka, M., et al., Endoplasmic reticulum remains continuous and undergoes sheet‐to‐tubule
transformation during cell division in mammalian cells. The Journal of cell biology, 2007.
179(5): p. 895‐909.
23. Benyamini, P., P. Webster, and D.I. Meyer, Knockdown of p180 eliminates the terminal
differentiation of a secretory cell line. Molecular biology of the cell, 2009. 20(2): p. 732‐44.
24. Orso, G., et al., Homotypic fusion of ER membranes requires the dynamin‐like GTPase
atlastin. Nature, 2009. 460(7258): p. 978‐83.
25. Hu, J., et al., A class of dynamin‐like GTPases involved in the generation of the tubular ER
network. Cell, 2009. 138(3): p. 549‐61.
26. Palade, G.E., A small particulate component of the cytoplasm. The Journal of biophysical and
biochemical cytology, 1955. 1(1): p. 59‐68.
27. Palade, G.E., The endoplasmic reticulum. The Journal of biophysical and biochemical
cytology, 1956. 2(4 Suppl): p. 85‐98.
28. Palade, G.E., Intracisternal granules in the exocrine cells of the pancreas. The Journal of
biophysical and biochemical cytology, 1956. 2(4): p. 417‐22.
29. Palade, G.E. and P. Siekevitz, Pancreatic microsomes; an integrated morphological and
biochemical study. The Journal of biophysical and biochemical cytology, 1956. 2(6): p. 671‐
90.
30. Palade, G.E. and P. Siekevitz, Liver microsomes; an integrated morphological and biochemical
study. The Journal of biophysical and biochemical cytology, 1956. 2(2): p. 171‐200.
31. Deshaies, R.J. and R. Schekman, A yeast mutant defective at an early stage in import of
secretory protein precursors into the endoplasmic reticulum. The Journal of cell biology,
1987. 105(2): p. 633‐45.
32. Krieg, U.C., A.E. Johnson, and P. Walter, Protein translocation across the endoplasmic
reticulum membrane: identification by photocross‐linking of a 39‐kD integral membrane
glycoprotein as part of a putative translocation tunnel. The Journal of cell biology, 1989.
109(5): p. 2033‐43.
33. Wiedmann, M., et al., Photocrosslinking demonstrates proximity of a 34 kDa membrane
protein to different portions of preprolactin during translocation through the endoplasmic
reticulum. FEBS letters, 1989. 257(2): p. 263‐8.
34. Gorlich, D., et al., A protein of the endoplasmic reticulum involved early in polypeptide
translocation. Nature, 1992. 357(6373): p. 47‐52.
35. Gorlich, D., et al., A mammalian homolog of SEC61p and SECYp is associated with ribosomes
and nascent polypeptides during translocation. Cell, 1992. 71(3): p. 489‐503.
36. Gorlich, D. and T.A. Rapoport, Protein translocation into proteoliposomes reconstituted from
purified components of the endoplasmic reticulum membrane. Cell, 1993. 75(4): p. 615‐30.
37. Do, H., et al., The cotranslational integration of membrane proteins into the phospholipid
bilayer is a multistep process. Cell, 1996. 85(3): p. 369‐78.
38. Martoglio, B., et al., The protein‐conducting channel in the membrane of the endoplasmic
reticulum is open laterally toward the lipid bilayer. Cell, 1995. 81(2): p. 207‐14.
39. Mothes, W., et al., Molecular mechanism of membrane protein integration into the
endoplasmic reticulum. Cell, 1997. 89(4): p. 523‐33.
40. Ellgaard, L. and A. Helenius, Quality control in the endoplasmic reticulum. Nature reviews.
Molecular cell biology, 2003. 4(3): p. 181‐91.

63
41. Brodsky, J.L. and W.R. Skach, Protein folding and quality control in the endoplasmic
reticulum: Recent lessons from yeast and mammalian cell systems. Current opinion in cell
biology, 2011. 23(4): p. 464‐75.
42. Brown, D.A. and R.D. Simoni, Biogenesis of 3‐hydroxy‐3‐methylglutaryl‐coenzyme A
reductase, an integral glycoprotein of the endoplasmic reticulum. Proceedings of the
National Academy of Sciences of the United States of America, 1984. 81(6): p. 1674‐8.
43. Skalnik, D.G., et al., The membrane domain of 3‐hydroxy‐3‐methylglutaryl‐coenzyme A
reductase confers endoplasmic reticulum localization and sterol‐regulated degradation onto
beta‐galactosidase. The Journal of biological chemistry, 1988. 263(14): p. 6836‐41.
44. Gil, G., et al., Membrane‐bound domain of HMG CoA reductase is required for sterol‐
enhanced degradation of the enzyme. Cell, 1985. 41(1): p. 249‐58.
45. Bannykh, S.I., T. Rowe, and W.E. Balch, The organization of endoplasmic reticulum export
complexes. J Cell Biol, 1996. 135(1): p. 19‐35.
46. Tartakoff, A.M., Temperature and energy dependence of secretory protein transport in the
exocrine pancreas. The EMBO journal, 1986. 5(7): p. 1477‐82.
47. Lotti, L.V., et al., Morphological analysis of the transfer of VSV ts‐045 G glycoprotein from the
endoplasmic reticulum to the intermediate compartment in vero cells. Experimental cell
research, 1996. 227(2): p. 323‐31.
48. Mezzacasa, A. and A. Helenius, The transitional ER defines a boundary for quality control in
the secretion of tsO45 VSV glycoprotein. Traffic, 2002. 3(11): p. 833‐49.
49. Saraste, J. and E. Kuismanen, Pre‐ and post‐Golgi vacuoles operate in the transport of Semliki
Forest virus membrane glycoproteins to the cell surface. Cell, 1984. 38(2): p. 535‐49.
50. Aridor, M., et al., Sequential coupling between COPII and COPI vesicle coats in endoplasmic
reticulum to Golgi transport. J Cell Biol, 1995. 131(4): p. 875‐93.
51. Peter, F., et al., Beta‐COP is essential for transport of protein from the endoplasmic reticulum
to the Golgi in vitro. J Cell Biol, 1993. 122(6): p. 1155‐67.
52. Scales, S.J., R. Pepperkok, and T.E. Kreis, Visualization of ER‐to‐Golgi transport in living cells
reveals a sequential mode of action for COPII and COPI. Cell, 1997. 90(6): p. 1137‐48.
53. Yu, S., et al., mBet3p is required for homotypic COPII vesicle tethering in mammalian cells.
The Journal of cell biology, 2006. 174(3): p. 359‐68.
54. Cai, H., et al., TRAPPI tethers COPII vesicles by binding the coat subunit Sec23. Nature, 2007.
445(7130): p. 941‐4.
55. Lord, C., et al., Sequential interactions with Sec23 control the direction of vesicle traffic.
Nature, 2011. 473(7346): p. 181‐6.
56. Jahn, R. and R.H. Scheller, SNAREs‐‐engines for membrane fusion. Nature reviews. Molecular
cell biology, 2006. 7(9): p. 631‐43.
57. Zhang, B., et al., Bleeding due to disruption of a cargo‐specific ER‐to‐Golgi transport complex.
Nat Genet, 2003. 34(2): p. 220‐5.
58. Nichols, W.C., et al., Mutations in the ER‐Golgi intermediate compartment protein ERGIC‐53
cause combined deficiency of coagulation factors V and VIII. Cell, 1998. 93(1): p. 61‐70.
59. Zheng, C., et al., EF‐hand domains of MCFD2 mediate interactions with both LMAN1 and
coagulation factor V or VIII. Blood, 2010. 115(5): p. 1081‐7.
60. Itin, C., et al., Recycling of the endoplasmic reticulum/Golgi intermediate compartment
protein ERGIC‐53 in the secretory pathway. Biochem Soc Trans, 1995. 23(3): p. 541‐4.
61. Schweizer, A., et al., Identification, by a monoclonal antibody, of a 53‐kD protein associated
with a tubulo‐vesicular compartment at the cis‐side of the Golgi apparatus. J Cell Biol, 1988.
107(5): p. 1643‐53.
62. Appenzeller, C., et al., The lectin ERGIC‐53 is a cargo transport receptor for glycoproteins. Nat
Cell Biol, 1999. 1(6): p. 330‐4.

64
63. Kappeler, F., et al., The recycling of ERGIC‐53 in the early secretory pathway. ERGIC‐53
carries a cytosolic endoplasmic reticulum‐exit determinant interacting with COPII. J Biol
Chem, 1997. 272(50): p. 31801‐8.
64. Nufer, O., et al., ER export of ERGIC‐53 is controlled by cooperation of targeting determinants
in all three of its domains. Journal of cell science, 2003. 116(Pt 21): p. 4429‐40.
65. Itin, C., R. Schindler, and H.P. Hauri, Targeting of protein ERGIC‐53 to the ER/ERGIC/cis‐Golgi
recycling pathway. J Cell Biol, 1995. 131(1): p. 57‐67.
66. Appenzeller‐Herzog, C., et al., pH‐induced conversion of the transport lectin ERGIC‐53
triggers glycoprotein release. J Biol Chem, 2004. 279(13): p. 12943‐50.
67. Bisel, B., et al., ERK regulates Golgi and centrosome orientation towards the leading edge
through GRASP65. J Cell Biol, 2008. 182(5): p. 837‐43.
68. Stinchcombe, J.C., et al., Centrosome polarization delivers secretory granules to the
immunological synapse. Nature, 2006. 443(7110): p. 462‐5.
69. Horton, A.C., et al., Polarized secretory trafficking directs cargo for asymmetric dendrite
growth and morphogenesis. Neuron, 2005. 48(5): p. 757‐71.
70. Vasiliev, J.M., et al., Effect of colcemid on the locomotory behaviour of fibroblasts. Journal of
embryology and experimental morphology, 1970. 24(3): p. 625‐40.
71. Yadav, S., S. Puri, and A.D. Linstedt, A primary role for Golgi positioning in directed secretion,
cell polarity, and wound healing. Molecular biology of the cell, 2009. 20(6): p. 1728‐36.
72. Hurtado, L., et al., Disconnecting the Golgi ribbon from the centrosome prevents directional
cell migration and ciliogenesis. The Journal of cell biology, 2011. 193(5): p. 917‐33.
73. Wilson, C., et al., The Golgi apparatus: an organelle with multiple complex functions.
Biochem J, 2011. 433(1): p. 1‐9.
74. Clermont, Y., A. Rambourg, and L. Hermo, Trans‐Golgi network (TGN) of different cell types:
three‐dimensional structural characteristics and variability. Anat Rec, 1995. 242(3): p. 289‐
301.
75. Ladinsky, M.S., et al., Golgi structure in three dimensions: functional insights from the normal
rat kidney cell. J Cell Biol, 1999. 144(6): p. 1135‐49.
76. Marsh, B.J., et al., Structural evidence for multiple transport mechanisms through the Golgi in
the pancreatic beta‐cell line, HIT‐T15. Biochemical Society transactions, 2001. 29(Pt 4): p.
461‐7.
77. Mogelsvang, S., et al., Predicting function from structure: 3D structure studies of the
mammalian Golgi complex. Traffic, 2004. 5(5): p. 338‐45.
78. Bonifacino, J.S. and L.M. Traub, Signals for sorting of transmembrane proteins to endosomes
and lysosomes. Annual review of biochemistry, 2003. 72: p. 395‐447.
79. Stamnes, M.A. and J.E. Rothman, The binding of AP‐1 clathrin adaptor particles to Golgi
membranes requires ADP‐ribosylation factor, a small GTP‐binding protein. Cell, 1993. 73(5):
p. 999‐1005.
80. Traub, L.M., J.A. Ostrom, and S. Kornfeld, Biochemical dissection of AP‐1 recruitment onto
Golgi membranes. The Journal of cell biology, 1993. 123(3): p. 561‐73.
81. Boman, A.L., et al., A family of ADP‐ribosylation factor effectors that can alter membrane
transport through the trans‐Golgi. Molecular biology of the cell, 2000. 11(4): p. 1241‐55.
82. Dell'Angelica, E.C., et al., GGAs: a family of ADP ribosylation factor‐binding proteins related
to adaptors and associated with the Golgi complex. The Journal of cell biology, 2000. 149(1):
p. 81‐94.
83. Hirst, J., et al., A family of proteins with gamma‐adaptin and VHS domains that facilitate
trafficking between the trans‐Golgi network and the vacuole/lysosome. The Journal of cell
biology, 2000. 149(1): p. 67‐80.
84. Bilodeau, P.S., et al., The GAT domains of clathrin‐associated GGA proteins have two
ubiquitin binding motifs. The Journal of biological chemistry, 2004. 279(52): p. 54808‐16.

65
85. Scott, P.M., et al., GGA proteins bind ubiquitin to facilitate sorting at the trans‐Golgi network.
Nature cell biology, 2004. 6(3): p. 252‐9.
86. Puertollano, R. and J.S. Bonifacino, Interactions of GGA3 with the ubiquitin sorting
machinery. Nature cell biology, 2004. 6(3): p. 244‐51.
87. Ladinsky, M.S., et al., Structure of the Golgi and distribution of reporter molecules at 20
degrees C reveals the complexity of the exit compartments. Molecular biology of the cell,
2002. 13(8): p. 2810‐25.
88. Glick, B.S. and A. Nakano, Membrane traffic within the Golgi apparatus. Annu Rev Cell Dev
Biol, 2009. 25: p. 113‐32.
89. Orci, L., B.S. Glick, and J.E. Rothman, A new type of coated vesicular carrier that appears not
to contain clathrin: its possible role in protein transport within the Golgi stack. Cell, 1986.
46(2): p. 171‐84.
90. Balch, W.E., B.S. Glick, and J.E. Rothman, Sequential intermediates in the pathway of
intercompartmental transport in a cell‐free system. Cell, 1984. 39(3 Pt 2): p. 525‐36.
91. Pelham, H.R. and J.E. Rothman, The debate about transport in the Golgi‐‐two sides of the
same coin? Cell, 2000. 102(6): p. 713‐9.
92. Franke, W.W., et al., Synthesis and turnover of membrane proteins in rat liver: an
examination of the membrane flow hypothesis. Z Naturforsch B, 1971. 26(10): p. 1031‐9.
93. Mironov, A.A., P. Weidman, and A. Luini, Variations on the intracellular transport theme:
maturing cisternae and trafficking tubules. J Cell Biol, 1997. 138(3): p. 481‐4.
94. Glick, B.S. and V. Malhotra, The curious status of the Golgi apparatus. Cell, 1998. 95(7): p.
883‐9.
95. Mironov, A.A., et al., Small cargo proteins and large aggregates can traverse the Golgi by a
common mechanism without leaving the lumen of cisternae. J Cell Biol, 2001. 155(7): p.
1225‐38.
96. Presley, J.F., et al., Dissection of COPI and Arf1 dynamics in vivo and role in Golgi membrane
transport. Nature, 2002. 417(6885): p. 187‐93.
97. Letourneur, F., et al., Coatomer is essential for retrieval of dilysine‐tagged proteins to the
endoplasmic reticulum. Cell, 1994. 79(7): p. 1199‐207.
98. Gommel, D.U., et al., Recruitment to Golgi membranes of ADP‐ribosylation factor 1 is
mediated by the cytoplasmic domain of p23. The EMBO journal, 2001. 20(23): p. 6751‐60.
99. Majoul, I., et al., KDEL‐cargo regulates interactions between proteins involved in COPI vesicle
traffic: measurements in living cells using FRET. Developmental cell, 2001. 1(1): p. 139‐53.
100. Contreras, I., E. Ortiz‐Zapater, and F. Aniento, Sorting signals in the cytosolic tail of
membrane proteins involved in the interaction with plant ARF1 and coatomer. The Plant
journal : for cell and molecular biology, 2004. 38(4): p. 685‐98.
101. Reinhard, C., et al., Receptor‐induced polymerization of coatomer. Proceedings of the
National Academy of Sciences of the United States of America, 1999. 96(4): p. 1224‐8.
102. Langer, J.D., et al., A conformational change in the alpha‐subunit of coatomer induced by
ligand binding to gamma‐COP revealed by single‐pair FRET. Traffic, 2008. 9(4): p. 597‐607.
103. Kahn, R.A. and A.G. Gilman, ADP‐ribosylation of Gs promotes the dissociation of its alpha and
beta subunits. The Journal of biological chemistry, 1984. 259(10): p. 6235‐40.
104. Chardin, P., et al., A human exchange factor for ARF contains Sec7‐ and pleckstrin‐homology
domains. Nature, 1996. 384(6608): p. 481‐4.
105. Helms, J.B. and J.E. Rothman, Inhibition by brefeldin A of a Golgi membrane enzyme that
catalyses exchange of guanine nucleotide bound to ARF. Nature, 1992. 360(6402): p. 352‐4.
106. Donaldson, J.G., D. Finazzi, and R.D. Klausner, Brefeldin A inhibits Golgi membrane‐catalysed
exchange of guanine nucleotide onto ARF protein. Nature, 1992. 360(6402): p. 350‐2.
107. Antonny, B., et al., N‐terminal hydrophobic residues of the G‐protein ADP‐ribosylation factor‐
1 insert into membrane phospholipids upon GDP to GTP exchange. Biochemistry, 1997.
36(15): p. 4675‐84.

66
108. Franco, M., et al., Myristoylation‐facilitated binding of the G protein ARF1GDP to membrane
phospholipids is required for its activation by a soluble nucleotide exchange factor. The
Journal of biological chemistry, 1996. 271(3): p. 1573‐8.
109. Liu, Y., R.A. Kahn, and J.H. Prestegard, Structure and membrane interaction of myristoylated
ARF1. Structure, 2009. 17(1): p. 79‐87.
110. Antonny, B., et al., Membrane curvature and the control of GTP hydrolysis in Arf1 during
COPI vesicle formation. Biochemical Society transactions, 2005. 33(Pt 4): p. 619‐22.
111. Bigay, J., et al., ArfGAP1 responds to membrane curvature through the folding of a lipid
packing sensor motif. The EMBO journal, 2005. 24(13): p. 2244‐53.
112. Bigay, J., et al., Lipid packing sensed by ArfGAP1 couples COPI coat disassembly to membrane
bilayer curvature. Nature, 2003. 426(6966): p. 563‐6.
113. Serafini, T., et al., A coat subunit of Golgi‐derived non‐clathrin‐coated vesicles with homology
to the clathrin‐coated vesicle coat protein beta‐adaptin. Nature, 1991. 349(6306): p. 215‐20.
114. Waters, M.G., T. Serafini, and J.E. Rothman, 'Coatomer': a cytosolic protein complex
containing subunits of non‐clathrin‐coated Golgi transport vesicles. Nature, 1991. 349(6306):
p. 248‐51.
115. Duden, R., et al., Beta‐COP, a 110 kd protein associated with non‐clathrin‐coated vesicles and
the Golgi complex, shows homology to beta‐adaptin. Cell, 1991. 64(3): p. 649‐65.
116. Stenbeck, G., et al., beta'‐COP, a novel subunit of coatomer. The EMBO journal, 1993. 12(7):
p. 2841‐5.
117. Harrison‐Lavoie, K.J., et al., A 102 kDa subunit of a Golgi‐associated particle has homology to
beta subunits of trimeric G proteins. The EMBO journal, 1993. 12(7): p. 2847‐53.
118. Hara‐Kuge, S., et al., En bloc incorporation of coatomer subunits during the assembly of COP‐
coated vesicles. The Journal of cell biology, 1994. 124(6): p. 883‐92.
119. Kuge, O., et al., zeta‐COP, a subunit of coatomer, is required for COP‐coated vesicle assembly.
The Journal of cell biology, 1993. 123(6 Pt 2): p. 1727‐34.
120. Lee, C. and J. Goldberg, Structure of coatomer cage proteins and the relationship among
COPI, COPII, and clathrin vesicle coats. Cell, 2010. 142(1): p. 123‐32.
121. Yu, X., M. Breitman, and J. Goldberg, A structure‐based mechanism for arf1‐dependent
recruitment of coatomer to membranes. Cell, 2012. 148(3): p. 530‐42.
122. Cosson, P. and F. Letourneur, Coatomer interaction with di‐lysine endoplasmic reticulum
retention motifs. Science, 1994. 263(5153): p. 1629‐31.
123. Jackson, M.R., T. Nilsson, and P.A. Peterson, Identification of a consensus motif for retention
of transmembrane proteins in the endoplasmic reticulum. The EMBO journal, 1990. 9(10): p.
3153‐62.
124. Dominguez, M., et al., gp25L/emp24/p24 protein family members of the cis‐Golgi network
bind both COP I and II coatomer. The Journal of cell biology, 1998. 140(4): p. 751‐65.
125. Gommel, D., et al., p24 and p23, the major transmembrane proteins of COPI‐coated
transport vesicles, form hetero‐oligomeric complexes and cycle between the organelles of the
early secretory pathway. FEBS letters, 1999. 447(2‐3): p. 179‐85.
126. Vincent, M.J., A.S. Martin, and R.W. Compans, Function of the KKXX motif in endoplasmic
reticulum retrieval of a transmembrane protein depends on the length and structure of the
cytoplasmic domain. The Journal of biological chemistry, 1998. 273(2): p. 950‐6.
127. Eugster, A., et al., The alpha‐ and beta'‐COP WD40 domains mediate cargo‐selective
interactions with distinct di‐lysine motifs. Molecular biology of the cell, 2004. 15(3): p. 1011‐
23.
128. Jackson, L.P., et al., Molecular basis for recognition of dilysine trafficking motifs by COPI. Dev
Cell, 2012. 23(6): p. 1255‐62.
129. Dean, N. and H.R. Pelham, Recycling of proteins from the Golgi compartment to the ER in
yeast. The Journal of cell biology, 1990. 111(2): p. 369‐77.

67
130. Pelham, H.R., Evidence that luminal ER proteins are sorted from secreted proteins in a post‐
ER compartment. The EMBO journal, 1988. 7(4): p. 913‐8.
131. Pelham, H.R., K.G. Hardwick, and M.J. Lewis, Sorting of soluble ER proteins in yeast. The
EMBO journal, 1988. 7(6): p. 1757‐62.
132. Wilson, D.W., M.J. Lewis, and H.R. Pelham, pH‐dependent binding of KDEL to its receptor in
vitro. The Journal of biological chemistry, 1993. 268(10): p. 7465‐8.
133. Lee, S.Y., et al., ARFGAP1 plays a central role in coupling COPI cargo sorting with vesicle
formation. The Journal of cell biology, 2005. 168(2): p. 281‐90.
134. Yang, J.S., et al., ARFGAP1 promotes the formation of COPI vesicles, suggesting function as a
component of the coat. The Journal of cell biology, 2002. 159(1): p. 69‐78.
135. Cukierman, E., et al., The ARF1 GTPase‐activating protein: zinc finger motif and Golgi
complex localization. Science, 1995. 270(5244): p. 1999‐2002.
136. Novick, P., C. Field, and R. Schekman, Identification of 23 complementation groups required
for post‐translational events in the yeast secretory pathway. Cell, 1980. 21(1): p. 205‐15.
137. Schekman, R., et al., Yeast secretory mutants: isolation and characterization. Methods
Enzymol, 1983. 96: p. 802‐15.
138. Ferro‐Novick, S., et al., Yeast secretory mutants that block the formation of active cell surface
enzymes. J Cell Biol, 1984. 98(1): p. 35‐43.
139. Kaiser, C.A. and R. Schekman, Distinct sets of SEC genes govern transport vesicle formation
and fusion early in the secretory pathway. Cell, 1990. 61(4): p. 723‐33.
140. Sato, K., COPII coat assembly and selective export from the endoplasmic reticulum. Journal of
biochemistry, 2004. 136(6): p. 755‐60.
141. Nakano, A., D. Brada, and R. Schekman, A membrane glycoprotein, Sec12p, required for
protein transport from the endoplasmic reticulum to the Golgi apparatus in yeast. J Cell Biol,
1988. 107(3): p. 851‐63.
142. Nishikawa, S. and A. Nakano, Identification of a gene required for membrane protein
retention in the early secretory pathway. Proc Natl Acad Sci U S A, 1993. 90(17): p. 8179‐83.
143. Sato, M., K. Sato, and A. Nakano, Endoplasmic reticulum localization of Sec12p is achieved by
two mechanisms: Rer1p‐dependent retrieval that requires the transmembrane domain and
Rer1p‐independent retention that involves the cytoplasmic domain. J Cell Biol, 1996. 134(2):
p. 279‐93.
144. d'Enfert, C., et al., Structural and functional dissection of a membrane glycoprotein required
for vesicle budding from the endoplasmic reticulum. Mol Cell Biol, 1991. 11(11): p. 5727‐34.
145. Nakano, A. and M. Muramatsu, A novel GTP‐binding protein, Sar1p, is involved in transport
from the endoplasmic reticulum to the Golgi apparatus. The Journal of cell biology, 1989.
109(6 Pt 1): p. 2677‐91.
146. d'Enfert, C., et al., Sec12p‐dependent membrane binding of the small GTP‐binding protein
Sar1p promotes formation of transport vesicles from the ER. J Cell Biol, 1991. 114(4): p. 663‐
70.
147. Barlowe, C. and R. Schekman, SEC12 encodes a guanine‐nucleotide‐exchange factor essential
for transport vesicle budding from the ER. Nature, 1993. 365(6444): p. 347‐9.
148. Weissman, J.T., H. Plutner, and W.E. Balch, The mammalian guanine nucleotide exchange
factor mSec12 is essential for activation of the Sar1 GTPase directing endoplasmic reticulum
export. Traffic, 2001. 2(7): p. 465‐75.
149. Lee, M.C., et al., Sar1p N‐terminal helix initiates membrane curvature and completes the
fission of a COPII vesicle. Cell, 2005. 122(4): p. 605‐17.
150. Bielli, A., et al., Regulation of Sar1 NH2 terminus by GTP binding and hydrolysis promotes
membrane deformation to control COPII vesicle fission. J Cell Biol, 2005. 171(6): p. 919‐24.
151. Bi, X., R.A. Corpina, and J. Goldberg, Structure of the Sec23/24‐Sar1 pre‐budding complex of
the COPII vesicle coat. Nature, 2002. 419(6904): p. 271‐7.

68
152. Huang, M., et al., Crystal structure of Sar1‐GDP at 1.7 A resolution and the role of the NH2
terminus in ER export. J Cell Biol, 2001. 155(6): p. 937‐48.
153. McMahon, C., et al., The Structure of Sec12 Implicates Potassium Ion Coordination in Sar1
Activation. J Biol Chem, 2012.
154. Spang, A., On vesicle formation and tethering in the ER‐Golgi shuttle. Curr Opin Cell Biol,
2009. 21(4): p. 531‐6.
155. Jones, B., et al., Mutations in a Sar1 GTPase of COPII vesicles are associated with lipid
absorption disorders. Nat Genet, 2003. 34(1): p. 29‐31.
156. Siddiqi, S.A., et al., COPII proteins are required for Golgi fusion but not for endoplasmic
reticulum budding of the pre‐chylomicron transport vesicle. J Cell Sci, 2003. 116(Pt 2): p. 415‐
27.
157. Aridor, M., et al., The Sar1 GTPase coordinates biosynthetic cargo selection with endoplasmic
reticulum export site assembly. J Cell Biol, 2001. 152(1): p. 213‐29.
158. Plutner, H., et al., Morphological analysis of protein transport from the ER to Golgi
membranes in digitonin‐permeabilized cells: role of the P58 containing compartment. The
Journal of cell biology, 1992. 119(5): p. 1097‐116.
159. Long, K.R., et al., Sar1 assembly regulates membrane constriction and ER export. J Cell Biol,
2010. 190(1): p. 115‐28.
160. Heinrich, V.V., S. Svetina, and B. Zeks, Nonaxisymmetric vesicle shapes in a generalized
bilayer‐couple model and the transition between oblate and prolate axisymmetric shapes.
Physical review. E, Statistical physics, plasmas, fluids, and related interdisciplinary topics,
1993. 48(4): p. 3112‐3123.
161. Antonny, B., et al., Dynamics of the COPII coat with GTP and stable analogues. Nat Cell Biol,
2001. 3(6): p. 531‐7.
162. Blumental‐Perry, A., et al., Phosphatidylinositol 4‐phosphate formation at ER exit sites
regulates ER export. Developmental cell, 2006. 11(5): p. 671‐82.
163. Saito, K., et al., TANGO1 facilitates cargo loading at endoplasmic reticulum exit sites. Cell,
2009. 136(5): p. 891‐902.
164. Venditti, R., et al., Sedlin controls the ER export of procollagen by regulating the Sar1 cycle.
Science, 2012. 337(6102): p. 1668‐72.
165. Saito‐Nakano, Y. and A. Nakano, Sed4p functions as a positive regulator of Sar1p probably
through inhibition of the GTPase activation by Sec23p. Genes to cells : devoted to molecular
& cellular mechanisms, 2000. 5(12): p. 1039‐48.
166. Kodera, C., et al., Sed4p stimulates Sar1p GTP hydrolysis and promotes limited coat
disassembly. Traffic, 2011. 12(5): p. 591‐9.
167. Barlowe, C., et al., COPII: a membrane coat formed by Sec proteins that drive vesicle budding
from the endoplasmic reticulum. Cell, 1994. 77(6): p. 895‐907.
168. Kuehn, M.J., J.M. Herrmann, and R. Schekman, COPII‐cargo interactions direct protein
sorting into ER‐derived transport vesicles. Nature, 1998. 391(6663): p. 187‐90.
169. Lederkremer, G.Z., et al., Structure of the Sec23p/24p and Sec13p/31p complexes of COPII.
Proc Natl Acad Sci U S A, 2001. 98(19): p. 10704‐9.
170. Bickford, L.C., E. Mossessova, and J. Goldberg, A structural view of the COPII vesicle coat.
Current opinion in structural biology, 2004. 14(2): p. 147‐53.
171. Peng, R., et al., Specific interaction of the yeast cis‐Golgi syntaxin Sed5p and the coat protein
complex II component Sec24p of endoplasmic reticulum‐derived transport vesicles.
Proceedings of the National Academy of Sciences of the United States of America, 1999.
96(7): p. 3751‐6.
172. Mossessova, E., L.C. Bickford, and J. Goldberg, SNARE selectivity of the COPII coat. Cell, 2003.
114(4): p. 483‐95.
173. Mancias, J.D. and J. Goldberg, The transport signal on Sec22 for packaging into COPII‐coated
vesicles is a conformational epitope. Mol Cell, 2007. 26(3): p. 403‐14.

69
174. Nishimura, N. and W.E. Balch, A di‐acidic signal required for selective export from the
endoplasmic reticulum. Science, 1997. 277(5325): p. 556‐8.
175. Miller, E.A., et al., Multiple cargo binding sites on the COPII subunit Sec24p ensure capture of
diverse membrane proteins into transport vesicles. Cell, 2003. 114(4): p. 497‐509.
176. Kung, L.F., et al., Sec24p and Sec16p cooperate to regulate the GTP cycle of the COPII coat.
The EMBO journal, 2011. 31(4): p. 1014‐27.
177. Fath, S., et al., Structure and organization of coat proteins in the COPII cage. Cell, 2007.
129(7): p. 1325‐36.
178. Brohawn, S.G., et al., Structural evidence for common ancestry of the nuclear pore complex
and vesicle coats. Science, 2008. 322(5906): p. 1369‐73.
179. Stagg, S.M., P. LaPointe, and W.E. Balch, Structural design of cage and coat scaffolds that
direct membrane traffic. Current opinion in structural biology, 2007. 17(2): p. 221‐8.
180. Stagg, S.M., et al., Structural basis for cargo regulation of COPII coat assembly. Cell, 2008.
134(3): p. 474‐84.
181. Bi, X., J.D. Mancias, and J. Goldberg, Insights into COPII coat nucleation from the structure of
Sec23.Sar1 complexed with the active fragment of Sec31. Developmental cell, 2007. 13(5): p.
635‐45.
182. Stagg, S.M., et al., Structure of the Sec13/31 COPII coat cage. Nature, 2006. 439(7073): p.
234‐8.
183. Gurkan, C., et al., The COPII cage: unifying principles of vesicle coat assembly. Nat Rev Mol
Cell Biol, 2006. 7(10): p. 727‐38.
184. Matsuoka, K., et al., COPII‐coated vesicle formation reconstituted with purified coat proteins
and chemically defined liposomes. Cell, 1998. 93(2): p. 263‐75.
185. Supek, F., et al., Sec16p potentiates the action of COPII proteins to bud transport vesicles. J
Cell Biol, 2002. 158(6): p. 1029‐38.
186. Forster, R., et al., Secretory cargo regulates the turnover of COPII subunits at single ER exit
sites. Curr Biol, 2006. 16(2): p. 173‐9.
187. Yamasaki, A., et al., The Ca2+‐binding protein ALG‐2 is recruited to endoplasmic reticulum
exit sites by Sec31A and stabilizes the localization of Sec31A. Mol Biol Cell, 2006. 17(11): p.
4876‐87.
188. Bentley, M., et al., Vesicular calcium regulates coat retention, fusogenicity, and size of pre‐
Golgi intermediates. Mol Biol Cell, 2010. 21(6): p. 1033‐46.
189. Shibata, H., et al., The ALG‐2 binding site in Sec31A influences the retention kinetics of
Sec31A at the endoplasmic reticulum exit sites as revealed by live‐cell time‐lapse imaging.
Biosci Biotechnol Biochem, 2010. 74(9): p. 1819‐26.
190. Rismanchi, N., R. Puertollano, and C. Blackstone, STAM adaptor proteins interact with COPII
complexes and function in ER‐to‐Golgi trafficking. Traffic, 2009. 10(2): p. 201‐17.
191. Miller, E., et al., Cargo selection into COPII vesicles is driven by the Sec24p subunit. EMBO J,
2002. 21(22): p. 6105‐13.
192. Giraudo, C.G. and H.J. Maccioni, Endoplasmic reticulum export of glycosyltransferases
depends on interaction of a cytoplasmic dibasic motif with Sar1. Molecular biology of the
cell, 2003. 14(9): p. 3753‐66.
193. Bannykh, S., et al., Regulated export of cargo from the endoplasmic reticulum of mammalian
cells. Cold Spring Harb Symp Quant Biol, 1995. 60: p. 127‐37.
194. Aridor, M., et al., Cargo selection by the COPII budding machinery during export from the ER.
J Cell Biol, 1998. 141(1): p. 61‐70.
195. Aridor, M., et al., Cargo can modulate COPII vesicle formation from the endoplasmic
reticulum. The Journal of biological chemistry, 1999. 274(7): p. 4389‐99.
196. Votsmeier, C. and D. Gallwitz, An acidic sequence of a putative yeast Golgi membrane
protein binds COPII and facilitates ER export. EMBO J, 2001. 20(23): p. 6742‐50.

70
197. Nufer, O., et al., Role of cytoplasmic C‐terminal amino acids of membrane proteins in ER
export. J Cell Sci, 2002. 115(Pt 3): p. 619‐28.
198. Pagano, A., et al., Sec24 proteins and sorting at the endoplasmic reticulum. J Biol Chem,
1999. 274(12): p. 7833‐40.
199. Wendeler, M.W., J.P. Paccaud, and H.P. Hauri, Role of Sec24 isoforms in selective export of
membrane proteins from the endoplasmic reticulum. EMBO Rep, 2007. 8(3): p. 258‐64.
200. Tang, B.L., et al., A family of mammalian proteins homologous to yeast Sec24p. Biochem
Biophys Res Commun, 1999. 258(3): p. 679‐84.
201. Stankewich, M.C., P.R. Stabach, and J.S. Morrow, Human Sec31B: a family of new
mammalian orthologues of yeast Sec31p that associate with the COPII coat. J Cell Sci, 2006.
119(Pt 5): p. 958‐69.
202. Tang, B.L., et al., Mammalian homologues of yeast sec31p. An ubiquitously expressed form is
localized to endoplasmic reticulum (ER) exit sites and is essential for ER‐Golgi transport. J Biol
Chem, 2000. 275(18): p. 13597‐604.
203. Sevier, C.S., et al., Efficient export of the vesicular stomatitis virus G protein from the
endoplasmic reticulum requires a signal in the cytoplasmic tail that includes both tyrosine‐
based and di‐acidic motifs. Mol Biol Cell, 2000. 11(1): p. 13‐22.
204. Barlowe, C., Molecular recognition of cargo by the COPII complex: a most accommodating
coat. Cell, 2003. 114(4): p. 395‐7.
205. Iodice, L., S. Sarnataro, and S. Bonatti, The carboxyl‐terminal valine is required for transport
of glycoprotein CD8 alpha from the endoplasmic reticulum to the intermediate compartment.
The Journal of biological chemistry, 2001. 276(31): p. 28920‐6.
206. Rothman, J.E. and F.T. Wieland, Protein sorting by transport vesicles. Science, 1996.
272(5259): p. 227‐34.
207. Denecke, J., J. Botterman, and R. Deblaere, Protein secretion in plant cells can occur via a
default pathway. Plant Cell, 1990. 2(1): p. 51‐9.
208. Belden, W.J. and C. Barlowe, Role of Erv29p in collecting soluble secretory proteins into ER‐
derived transport vesicles. Science, 2001. 294(5546): p. 1528‐31.
209. Otte, S., et al., Erv41p and Erv46p: new components of COPII vesicles involved in transport
between the ER and Golgi complex. J Cell Biol, 2001. 152(3): p. 503‐18.
210. Caldwell, S.R., K.J. Hill, and A.A. Cooper, Degradation of endoplasmic reticulum (ER) quality
control substrates requires transport between the ER and Golgi. J Biol Chem, 2001. 276(26):
p. 23296‐303.
211. Dancourt, J. and C. Barlowe, Protein sorting receptors in the early secretory pathway. Annu
Rev Biochem, 2010. 79: p. 777‐802.
212. Wiseman, R.L., et al., Protein energetics in maturation of the early secretory pathway. Curr
Opin Cell Biol, 2007. 19(4): p. 359‐67.
213. Lang, M.R., et al., Secretory COPII coat component Sec23a is essential for craniofacial
chondrocyte maturation. Nat Genet, 2006. 38(10): p. 1198‐203.
214. Townley, A.K., et al., Efficient coupling of Sec23‐Sec24 to Sec13‐Sec31 drives COPII‐
dependent collagen secretion and is essential for normal craniofacial development. J Cell Sci,
2008. 121(Pt 18): p. 3025‐34.
215. Boyadjiev, S.A., et al., Cranio‐lenticulo‐sutural dysplasia associated with defects in collagen
secretion. Clin Genet, 2011. 80(2): p. 169‐76.
216. Fromme, J.C., et al., The genetic basis of a craniofacial disease provides insight into COPII
coat assembly. Developmental cell, 2007. 13(5): p. 623‐34.
217. Kim, S.D., et al., The SEC23‐SEC31 interface plays a critical role for export of procollagen from
the endoplasmic reticulum. The Journal of biological chemistry, 2012.
218. Charcosset, M., et al., Anderson or chylomicron retention disease: molecular impact of five
mutations in the SAR1B gene on the structure and the functionality of Sar1b protein.
Molecular genetics and metabolism, 2008. 93(1): p. 74‐84.

71
219. Siddiqi, S., S.A. Siddiqi, and C.M. Mansbach, 2nd, Sec24C is required for docking the
prechylomicron transport vesicle with the Golgi. J Lipid Res, 2010. 51(5): p. 1093‐100.
220. Schwarz, K., et al., Mutations affecting the secretory COPII coat component SEC23B cause
congenital dyserythropoietic anemia type II. Nature genetics, 2009. 41(8): p. 936‐40.
221. Wansleeben, C., et al., Planar cell polarity defects and defective Vangl2 trafficking in mutants
for the COPII gene Sec24b. Development, 2010. 137(7): p. 1067‐73.
222. Everett, B., Zhang, B., Vasievich, M., Adams, E., Khoriaty, R., Chen, X‐W. & Ginsburg, D. ,
COPII components SEC23A and SEC23B are required for normal murine development
(Poster), 2011, ASCB Annual Meeting 2011: Denver, CO.
223. Adams, E., Baines, A. & Ginsburg, D. , The Role of Mammalian COPII Component SEC24D
(Poster), 2011, ASCB Annual Meeting 2011: Denver, CO.
224. Singer, S.J. and G.L. Nicolson, The fluid mosaic model of the structure of cell membranes.
Science, 1972. 175(4023): p. 720‐31.
225. Hermansson, M., K. Hokynar, and P. Somerharju, Mechanisms of glycerophospholipid
homeostasis in mammalian cells. Progress in lipid research, 2011. 50(3): p. 240‐57.
226. van Meer, G., D.R. Voelker, and G.W. Feigenson, Membrane lipids: where they are and how
they behave. Nat Rev Mol Cell Biol, 2008. 9(2): p. 112‐24.
227. van Meer, G. and A.I. de Kroon, Lipid map of the mammalian cell. J Cell Sci, 2011. 124(Pt 1):
p. 5‐8.
228. Jelsema, C.L. and D.J. Morre, Distribution of phospholipid biosynthetic enzymes among cell
components of rat liver. J Biol Chem, 1978. 253(21): p. 7960‐71.
229. Bell, R.M., L.M. Ballas, and R.A. Coleman, Lipid topogenesis. J Lipid Res, 1981. 22(3): p. 391‐
403.
230. Sprong, H., et al., UDP‐galactose:ceramide galactosyltransferase is a class I integral
membrane protein of the endoplasmic reticulum. J Biol Chem, 1998. 273(40): p. 25880‐8.
231. Futerman, A.H. and H. Riezman, The ins and outs of sphingolipid synthesis. Trends Cell Biol,
2005. 15(6): p. 312‐8.
232. Henneberry, A.L., M.M. Wright, and C.R. McMaster, The major sites of cellular phospholipid
synthesis and molecular determinants of Fatty Acid and lipid head group specificity. Mol Biol
Cell, 2002. 13(9): p. 3148‐61.
233. Voelker, D.R., Bridging gaps in phospholipid transport. Trends Biochem Sci, 2005. 30(7): p.
396‐404.
234. Kennedy, E.P., Metabolism of lipides. Annual review of biochemistry, 1957. 26: p. 119‐48.
235. Bell, R.M. and R.A. Coleman, Enzymes of glycerolipid synthesis in eukaryotes. Annual review
of biochemistry, 1980. 49: p. 459‐87.
236. Bishop, W.R. and R.M. Bell, Assembly of phospholipids into cellular membranes: biosynthesis,
transmembrane movement and intracellular translocation. Annual review of cell biology,
1988. 4: p. 579‐610.
237. Kent, C., Eukaryotic phospholipid biosynthesis. Annual review of biochemistry, 1995. 64: p.
315‐43.
238. Imai, A. and M.C. Gershengorn, Independent phosphatidylinositol synthesis in pituitary
plasma membrane and endoplasmic reticulum. Nature, 1987. 325(6106): p. 726‐8.
239. Galvao, C. and J.A. Shayman, The phosphatidylinositol synthase of proximal tubule cells.
Biochim Biophys Acta, 1990. 1044(1): p. 34‐42.
240. Anderson, R.A. and V.T. Marchesi, Regulation of the association of membrane skeletal
protein 4.1 with glycophorin by a polyphosphoinositide. Nature, 1985. 318(6043): p. 295‐8.
241. Burn, P., Phosphatidylinositol cycle and its possible involvement in the regulation of
cytoskeleton‐membrane interactions. Journal of cellular biochemistry, 1988. 36(1): p. 15‐24.
242. Martin, T.F., Phosphoinositide lipids as signaling molecules: common themes for signal
transduction, cytoskeletal regulation, and membrane trafficking. Annual review of cell and
developmental biology, 1998. 14: p. 231‐64.

72
243. Lev, S., Non‐vesicular lipid transport by lipid‐transfer proteins and beyond. Nature reviews.
Molecular cell biology, 2010. 11(10): p. 739‐50.
244. Bishop, W.R. and R.M. Bell, Assembly of the endoplasmic reticulum phospholipid bilayer: the
phosphatidylcholine transporter. Cell, 1985. 42(1): p. 51‐60.
245. Kawashima, Y. and R.M. Bell, Assembly of the endoplasmic reticulum phospholipid bilayer.
Transporters for phosphatidylcholine and metabolites. The Journal of biological chemistry,
1987. 262(34): p. 16495‐502.
246. Bevers, E.M., et al., Generation of prothrombin‐converting activity and the exposure of
phosphatidylserine at the outer surface of platelets. European journal of biochemistry / FEBS,
1982. 122(2): p. 429‐36.
247. Bitbol, M. and P.F. Devaux, Measurement of outward translocation of phospholipids across
human erythrocyte membrane. Proceedings of the National Academy of Sciences of the
United States of America, 1988. 85(18): p. 6783‐7.
248. Connor, J., et al., Bidirectional transbilayer movement of phospholipid analogs in human red
blood cells. Evidence for an ATP‐dependent and protein‐mediated process. The Journal of
biological chemistry, 1992. 267(27): p. 19412‐7.
249. Helmkamp, G.M., Jr., et al., Phospholipid exchange between membranes. Purification of
bovine brain proteins that preferentially catalyze the transfer of phosphatidylinositol. The
Journal of biological chemistry, 1974. 249(20): p. 6382‐9.
250. Crain, R.C. and D.B. Zilversmit, Net transfer of phospholipid by the nonspecific phospholipid
transfer proteins from bovine liver. Biochimica et biophysica acta, 1980. 620(1): p. 37‐48.
251. Crain, R.C. and D.B. Zilversmit, Two nonspecific phospholipid exchange proteins from beef
liver. I. Purification and characterization. Biochemistry, 1980. 19(7): p. 1433‐9.
252. Morton, R.E. and D.B. Zilversmit, Purification and characterization of lipid transfer protein(s)
from human lipoprotein‐deficient plasma. Journal of lipid research, 1982. 23(7): p. 1058‐67.
253. Young, W.W., Jr., M.S. Lutz, and W.A. Blackburn, Endogenous glycosphingolipids move to the
cell surface at a rate consistent with bulk flow estimates. J Biol Chem, 1992. 267(17): p.
12011‐5.
254. Gillon, A.D., C.F. Latham, and E.A. Miller, Vesicle‐mediated ER export of proteins and lipids.
Biochim Biophys Acta, 2012. 1821(8): p. 1040‐9.
255. Vance, J.E., E.J. Aasman, and R. Szarka, Brefeldin A does not inhibit the movement of
phosphatidylethanolamine from its sites for synthesis to the cell surface. J Biol Chem, 1991.
266(13): p. 8241‐7.
256. Kaplan, M.R. and R.D. Simoni, Intracellular transport of phosphatidylcholine to the plasma
membrane. J Cell Biol, 1985. 101(2): p. 441‐5.
257. Gnamusch, E., et al., Transport of phospholipids between subcellular membranes of wild‐type
yeast cells and of the phosphatidylinositol transfer protein‐deficient strain Saccharomyces
cerevisiae sec 14. Biochim Biophys Acta, 1992. 1111(1): p. 120‐6.
258. Kawano, M., et al., Efficient trafficking of ceramide from the endoplasmic reticulum to the
Golgi apparatus requires a VAMP‐associated protein‐interacting FFAT motif of CERT. The
Journal of biological chemistry, 2006. 281(40): p. 30279‐88.
259. Chandran, S. and C.E. Machamer, Acute perturbations in Golgi organization impact de novo
sphingomyelin synthesis. Traffic, 2008. 9(11): p. 1894‐904.
260. Perry, R.J. and N.D. Ridgway, Oxysterol‐binding protein and vesicle‐associated membrane
protein‐associated protein are required for sterol‐dependent activation of the ceramide
transport protein. Molecular biology of the cell, 2006. 17(6): p. 2604‐16.
261. Whatmore, J., et al., Resynthesis of phosphatidylinositol in permeabilized neutrophils
following phospholipase Cbeta activation: transport of the intermediate, phosphatidic acid,
from the plasma membrane to the endoplasmic reticulum for phosphatidylinositol
resynthesis is not dependent on soluble lipid carriers or vesicular transport. The Biochemical
journal, 1999. 341 ( Pt 2): p. 435‐44.

73
262. Franke, W.W. and J. Kartenbeck, Outer mitochondrial membrane continuous with
endoplasmic reticulum. Protoplasma, 1971. 73(1): p. 35‐41.
263. Ardail, D., F. Lerme, and P. Louisot, Involvement of contact sites in phosphatidylserine import
into liver mitochondria. J Biol Chem, 1991. 266(13): p. 7978‐81.
264. Vance, J.E., Phospholipid synthesis in a membrane fraction associated with mitochondria. J
Biol Chem, 1990. 265(13): p. 7248‐56.
265. Shiao, Y.J., G. Lupo, and J.E. Vance, Evidence that phosphatidylserine is imported into
mitochondria via a mitochondria‐associated membrane and that the majority of
mitochondrial phosphatidylethanolamine is derived from decarboxylation of
phosphatidylserine. J Biol Chem, 1995. 270(19): p. 11190‐8.
266. Achleitner, G., et al., Association between the endoplasmic reticulum and mitochondria of
yeast facilitates interorganelle transport of phospholipids through membrane contact. Eur J
Biochem, 1999. 264(2): p. 545‐53.
267. Gaigg, B., et al., Characterization of a microsomal subfraction associated with mitochondria
of the yeast, Saccharomyces cerevisiae. Involvement in synthesis and import of phospholipids
into mitochondria. Biochim Biophys Acta, 1995. 1234(2): p. 214‐20.
268. Pichler, H., et al., A subfraction of the yeast endoplasmic reticulum associates with the
plasma membrane and has a high capacity to synthesize lipids. Eur J Biochem, 2001. 268(8):
p. 2351‐61.
269. Frolov, V.A., A.V. Shnyrova, and J. Zimmerberg, Lipid polymorphisms and membrane shape.
Cold Spring Harb Perspect Biol, 2011. 3(11): p. a004747.
270. Lindblom, G. and G. Oradd, Lipid lateral diffusion and membrane heterogeneity. Biochim
Biophys Acta, 2009. 1788(1): p. 234‐44.
271. Aikawa, Y., et al., Involvement of PITPnm, a mammalian homologue of Drosophila rdgB, in
phosphoinositide synthesis on Golgi membranes. The Journal of biological chemistry, 1999.
274(29): p. 20569‐77.
272. Los, D.A. and N. Murata, Membrane fluidity and its roles in the perception of environmental
signals. Biochim Biophys Acta, 2004. 1666(1‐2): p. 142‐57.
273. Ipsen, J.H., et al., Phase equilibria in the phosphatidylcholine‐cholesterol system. Biochim
Biophys Acta, 1987. 905(1): p. 162‐72.
274. Brown, D.A. and E. London, Structure of detergent‐resistant membrane domains: does phase
separation occur in biological membranes? Biochem Biophys Res Commun, 1997. 240(1): p.
1‐7.
275. Mannock, D.A., et al., The effect of variations in phospholipid and sterol structure on the
nature of lipid‐sterol interactions in lipid bilayer model membranes. Chem Phys Lipids, 2010.
163(6): p. 403‐48.
276. Baumgart, T., S.T. Hess, and W.W. Webb, Imaging coexisting fluid domains in biomembrane
models coupling curvature and line tension. Nature, 2003. 425(6960): p. 821‐4.
277. Vind‐Kezunovic, D., et al., Line tension at lipid phase boundaries regulates formation of
membrane vesicles in living cells. Biochim Biophys Acta, 2008. 1778(11): p. 2480‐6.
278. Vance, D.E. and H. Van den Bosch, Cholesterol in the year 2000. Biochim Biophys Acta, 2000.
1529(1‐3): p. 1‐8.
279. Liscum, L., Cholesterol biosynthesis, in Biochemistry of Lipids, Lipoproteins and Membranes,
D.E.V. Vance, J.E., Editor. 2002, Elsevier Science B.V. p. 409‐431.
280. Brown, M.S. and J.L. Goldstein, Receptor‐mediated control of cholesterol metabolism.
Science, 1976. 191(4223): p. 150‐4.
281. Yu, J., D.A. Fischman, and T.L. Steck, Selective solubilization of proteins and phospholipids
from red blood cell membranes by nonionic detergents. J Supramol Struct, 1973. 1(3): p. 233‐
48.
282. Simons, K. and G. van Meer, Lipid sorting in epithelial cells. Biochemistry, 1988. 27(17): p.
6197‐202.

74
283. Brown, D.A. and J.K. Rose, Sorting of GPI‐anchored proteins to glycolipid‐enriched membrane
subdomains during transport to the apical cell surface. Cell, 1992. 68(3): p. 533‐44.
284. Colbeau, A., J. Nachbaur, and P.M. Vignais, Enzymic characterization and lipid composition of
rat liver subcellular membranes. Biochim Biophys Acta, 1971. 249(2): p. 462‐92.
285. Lange, Y., et al., Regulation of endoplasmic reticulum cholesterol by plasma membrane
cholesterol. J Lipid Res, 1999. 40(12): p. 2264‐70.
286. Runz, H., et al., Sterols regulate ER‐export dynamics of secretory cargo protein ts‐O45‐G.
EMBO J, 2006. 25(13): p. 2953‐65.
287. Muniz, M., P. Morsomme, and H. Riezman, Protein sorting upon exit from the endoplasmic
reticulum. Cell, 2001. 104(2): p. 313‐20.
288. Heino, S., et al., Dissecting the role of the golgi complex and lipid rafts in biosynthetic
transport of cholesterol to the cell surface. Proc Natl Acad Sci U S A, 2000. 97(15): p. 8375‐80.
289. Ridsdale, A., et al., Cholesterol is required for efficient endoplasmic reticulum‐to‐Golgi
transport of secretory membrane proteins. Mol Biol Cell, 2006. 17(4): p. 1593‐605.
290. Israelachvili, J.N., D.J. Mitchell, and B.W. Ninham, Theory of self‐assembly of lipid bilayers
and vesicles. Biochim Biophys Acta, 1977. 470(2): p. 185‐201.
291. McMahon, H.T. and J.L. Gallop, Membrane curvature and mechanisms of dynamic cell
membrane remodelling. Nature, 2005. 438(7068): p. 590‐6.
292. Kooijman, E.E., et al., Spontaneous curvature of phosphatidic acid and lysophosphatidic acid.
Biochemistry, 2005. 44(6): p. 2097‐102.
293. Hammond, K., et al., Characterisation of phosphatidylcholine/phosphatidylinositol sonicated
vesicles. Effects of phospholipid composition on vesicle size. Biochim Biophys Acta, 1984.
774(1): p. 19‐25.
294. Sebastian, T.T., et al., Phospholipid flippases: building asymmetric membranes and transport
vesicles. Biochim Biophys Acta, 2012. 1821(8): p. 1068‐77.
295. Chen, C.Y., et al., Role for Drs2p, a P‐type ATPase and potential aminophospholipid
translocase, in yeast late Golgi function. J Cell Biol, 1999. 147(6): p. 1223‐36.
296. Chen, B., et al., Endocytic sorting and recycling require membrane phosphatidylserine
asymmetry maintained by TAT‐1/CHAT‐1. PLoS Genet, 2010. 6(12): p. e1001235.
297. Bettache, N., et al., Mechanical constraint imposed on plasma membrane through transverse
phospholipid imbalance induces reversible actin polymerization via phosphoinositide 3‐kinase
activation. J Cell Sci, 2003. 116(Pt 11): p. 2277‐84.
298. Vale, R.D., The molecular motor toolbox for intracellular transport. Cell, 2003. 112(4): p. 467‐
80.
299. Zimmerberg, J. and M.M. Kozlov, How proteins produce cellular membrane curvature. Nat
Rev Mol Cell Biol, 2006. 7(1): p. 9‐19.
300. Chernomordik, L., M.M. Kozlov, and J. Zimmerberg, Lipids in biological membrane fusion. J
Membr Biol, 1995. 146(1): p. 1‐14.
301. Takei, K., et al., Tubular membrane invaginations coated by dynamin rings are induced by
GTP‐gamma S in nerve terminals. Nature, 1995. 374(6518): p. 186‐90.
302. Bodenmiller, B., et al., PhosphoPep‐‐a phosphoproteome resource for systems biology
research in Drosophila Kc167 cells. Molecular systems biology, 2007. 3: p. 139.
303. Farsad, K., et al., Generation of high curvature membranes mediated by direct endophilin
bilayer interactions. J Cell Biol, 2001. 155(2): p. 193‐200.
304. Fabbri, M., S. Bannykh, and W.E. Balch, Export of protein from the endoplasmic reticulum is
regulated by a diacylglycerol/phorbol ester binding protein. J Biol Chem, 1994. 269(43): p.
26848‐57.
305. Gschwendt, M., W. Kittstein, and F. Marks, Protein kinase C activation by phorbol esters: do
cysteine‐rich regions and pseudosubstrate motifs play a role? Trends Biochem Sci, 1991.
16(5): p. 167‐9.

75
306. Szule, J.A., N.L. Fuller, and R.P. Rand, The effects of acyl chain length and saturation of
diacylglycerols and phosphatidylcholines on membrane monolayer curvature. Biophys J,
2002. 83(2): p. 977‐84.
307. Schmidt, A., et al., Endophilin I mediates synaptic vesicle formation by transfer of
arachidonate to lysophosphatidic acid. Nature, 1999. 401(6749): p. 133‐41.
308. Bi, K., M.G. Roth, and N.T. Ktistakis, Phosphatidic acid formation by phospholipase D is
required for transport from the endoplasmic reticulum to the Golgi complex. Curr Biol, 1997.
7(5): p. 301‐7.
309. Nagaya, H., et al., Diacylglycerol kinase delta suppresses ER‐to‐Golgi traffic via its SAM and
PH domains. Molecular biology of the cell, 2002. 13(1): p. 302‐16.
310. Pathre, P., et al., Activation of phospholipase D by the small GTPase Sar1p is required to
support COPII assembly and ER export. EMBO J, 2003. 22(16): p. 4059‐69.
311. Brown, W.J., et al., The lysophospholipid acyltransferase antagonist CI‐976 inhibits a late
step in COPII vesicle budding. Traffic, 2008. 9(5): p. 786‐97.
312. Zhao, C., et al., Phospholipase D2‐generated phosphatidic acid couples EGFR stimulation to
Ras activation by Sos. Nat Cell Biol, 2007. 9(6): p. 706‐12.
313. Vance, J.E. and R. Steenbergen, Metabolism and functions of phosphatidylserine. Prog Lipid
Res, 2005. 44(4): p. 207‐34.
314. James, D.J., et al., Phosphatidylinositol 4,5‐bisphosphate regulates SNARE‐dependent
membrane fusion. J Cell Biol, 2008. 182(2): p. 355‐66.
315. Chernomordik, L.V. and J. Zimmerberg, Bending membranes to the task: structural
intermediates in bilayer fusion. Curr Opin Struct Biol, 1995. 5(4): p. 541‐7.
316. De Matteis, M.A. and A. Godi, PI‐loting membrane traffic. Nature cell biology, 2004. 6(6): p.
487‐92.
317. Hay, J.C., et al., ATP‐dependent inositide phosphorylation required for Ca(2+)‐activated
secretion. Nature, 1995. 374(6518): p. 173‐7.
318. Mayinger, P., Phosphoinositides and vesicular membrane traffic. Biochim Biophys Acta, 2012.
1821(8): p. 1104‐13.
319. Jost, M., et al., Phosphatidylinositol‐4,5‐bisphosphate is required for endocytic coated vesicle
formation. Current biology : CB, 1998. 8(25): p. 1399‐402.
320. Krauss, M., et al., ARF6 stimulates clathrin/AP‐2 recruitment to synaptic membranes by
activating phosphatidylinositol phosphate kinase type Igamma. The Journal of cell biology,
2003. 162(1): p. 113‐24.
321. McPherson, P.S., et al., A presynaptic inositol‐5‐phosphatase. Nature, 1996. 379(6563): p.
353‐7.
322. Rohde, G., D. Wenzel, and V. Haucke, A phosphatidylinositol (4,5)‐bisphosphate binding site
within mu2‐adaptin regulates clathrin‐mediated endocytosis. The Journal of cell biology,
2002. 158(2): p. 209‐14.
323. Gaidarov, I. and J.H. Keen, Phosphoinositide‐AP‐2 interactions required for targeting to
plasma membrane clathrin‐coated pits. The Journal of cell biology, 1999. 146(4): p. 755‐64.
324. Lemmon, M.A., Membrane recognition by phospholipid‐binding domains. Nat Rev Mol Cell
Biol, 2008. 9(2): p. 99‐111.
325. McMahon, H.T. and E. Boucrot, Molecular mechanism and physiological functions of
clathrin‐mediated endocytosis. Nat Rev Mol Cell Biol, 2011. 12(8): p. 517‐33.
326. Stowell, M.H., et al., Nucleotide‐dependent conformational changes in dynamin: evidence for
a mechanochemical molecular spring. Nat Cell Biol, 1999. 1(1): p. 27‐32.
327. Cremona, O., et al., Essential role of phosphoinositide metabolism in synaptic vesicle
recycling. Cell, 1999. 99(2): p. 179‐88.
328. Fernandez‐Borja, M., et al., Multivesicular body morphogenesis requires phosphatidyl‐
inositol 3‐kinase activity. Curr Biol, 1999. 9(1): p. 55‐8.

76
329. Burman, C. and N.T. Ktistakis, Regulation of autophagy by phosphatidylinositol 3‐phosphate.
FEBS Lett, 2010. 584(7): p. 1302‐12.
330. Grainger, D.L., et al., The emerging role of PtdIns5P: another signalling phosphoinositide
takes its place. Biochem Soc Trans, 2012. 40(1): p. 257‐61.
331. Sasaki, T., et al., The physiology of phosphoinositides. Biol Pharm Bull, 2007. 30(9): p. 1599‐
604.
332. Di Paolo, G. and P. De Camilli, Phosphoinositides in cell regulation and membrane dynamics.
Nature, 2006. 443(7112): p. 651‐7.
333. Allan, D., Mapping the lipid distribution in the membranes of BHK cells (mini‐review).
Molecular membrane biology, 1996. 13(2): p. 81‐4.
334. Op den Kamp, J.A., Lipid asymmetry in membranes. Annual review of biochemistry, 1979. 48:
p. 47‐71.
335. Bretscher, M.S., Asymmetrical lipid bilayer structure for biological membranes. Nature: New
biology, 1972. 236(61): p. 11‐2.
336. Wang, Y.J., et al., Phosphatidylinositol 4 phosphate regulates targeting of clathrin adaptor
AP‐1 complexes to the Golgi. Cell, 2003. 114(3): p. 299‐310.
337. Balla, A., et al., A plasma membrane pool of phosphatidylinositol 4‐phosphate is generated
by phosphatidylinositol 4‐kinase type‐III alpha: studies with the PH domains of the oxysterol
binding protein and FAPP1. Molecular biology of the cell, 2005. 16(3): p. 1282‐95.
338. Wong, K., R. Meyers dd, and L.C. Cantley, Subcellular locations of phosphatidylinositol 4‐
kinase isoforms. The Journal of biological chemistry, 1997. 272(20): p. 13236‐41.
339. Manford, A.G., et al., ER‐to‐Plasma Membrane Tethering Proteins Regulate Cell Signaling
and ER Morphology. Dev Cell, 2012. 23(6): p. 1129‐40.
340. Nakatsu, F., et al., PtdIns4P synthesis by PI4KIIIalpha at the plasma membrane and its impact
on plasma membrane identity. J Cell Biol, 2012. 199(6): p. 1003‐16.
341. Tahirovic, S., M. Schorr, and P. Mayinger, Regulation of intracellular phosphatidylinositol‐4‐
phosphate by the Sac1 lipid phosphatase. Traffic, 2005. 6(2): p. 116‐30.
342. Blagoveshchenskaya, A., et al., Integration of Golgi trafficking and growth factor signaling by
the lipid phosphatase SAC1. The Journal of cell biology, 2008. 180(4): p. 803‐12.
343. Stefan, C.J., et al., Osh proteins regulate phosphoinositide metabolism at ER‐plasma
membrane contact sites. Cell, 2011. 144(3): p. 389‐401.
344. Rohde, H.M., et al., The human phosphatidylinositol phosphatase SAC1 interacts with the
coatomer I complex. The Journal of biological chemistry, 2003. 278(52): p. 52689‐99.
345. Malecz, N., et al., Synaptojanin 2, a novel Rac1 effector that regulates clathrin‐mediated
endocytosis. Curr Biol, 2000. 10(21): p. 1383‐6.
346. Christoforidis, S., et al., Phosphatidylinositol‐3‐OH kinases are Rab5 effectors. Nat Cell Biol,
1999. 1(4): p. 249‐52.
347. Harlan, J.E., et al., Pleckstrin homology domains bind to phosphatidylinositol‐4,5‐
bisphosphate. Nature, 1994. 371(6493): p. 168‐70.
348. Lemmon, M.A. and K.M. Ferguson, Signal‐dependent membrane targeting by pleckstrin
homology (PH) domains. Biochem J, 2000. 350 Pt 1: p. 1‐18.
349. Kanai, F., et al., The PX domains of p47phox and p40phox bind to lipid products of PI(3)K. Nat
Cell Biol, 2001. 3(7): p. 675‐8.
350. Cheever, M.L., et al., Phox domain interaction with PtdIns(3)P targets the Vam7 t‐SNARE to
vacuole membranes. Nat Cell Biol, 2001. 3(7): p. 613‐8.
351. Gaullier, J.M., et al., FYVE fingers bind PtdIns(3)P. Nature, 1998. 394(6692): p. 432‐3.
352. Patki, V., et al., A functional PtdIns(3)P‐binding motif. Nature, 1998. 394(6692): p. 433‐4.
353. Niggli, V., et al., Identification of a phosphatidylinositol‐4,5‐bisphosphate‐binding domain in
the N‐terminal region of ezrin. FEBS Lett, 1995. 376(3): p. 172‐6.
354. Tani, K., et al., p125 is a novel mammalian Sec23p‐interacting protein with structural
similarity to phospholipid‐modifying proteins. J Biol Chem, 1999. 274(29): p. 20505‐12.

77
355. Higgs, H.N., et al., Cloning of a phosphatidic acid‐preferring phospholipase A1 from bovine
testis. J Biol Chem, 1998. 273(10): p. 5468‐77.
356. Ford, M.G., et al., Simultaneous binding of PtdIns(4,5)P2 and clathrin by AP180 in the
nucleation of clathrin lattices on membranes. Science, 2001. 291(5506): p. 1051‐5.
357. Santagata, S., et al., G‐protein signaling through tubby proteins. Science, 2001. 292(5524): p.
2041‐50.
358. Farhan, H., et al., Adaptation of endoplasmic reticulum exit sites to acute and chronic
increases in cargo load. EMBO J, 2008. 27(15): p. 2043‐54.
359. Lorente‐Rodriguez, A. and C. Barlowe, Requirement for Golgi‐localized PI(4)P in fusion of
COPII vesicles with Golgi compartments. Mol Biol Cell, 2011. 22(2): p. 216‐29.
360. Carvou, N., et al., Phosphatidylinositol‐ and phosphatidylcholine‐transfer activity of PITPbeta
is essential for COPI‐mediated retrograde transport from the Golgi to the endoplasmic
reticulum. Journal of cell science, 2010. 123(Pt 8): p. 1262‐73.
361. Barylko, B., et al., A novel family of phosphatidylinositol 4‐kinases conserved from yeast to
humans. The Journal of biological chemistry, 2001. 276(11): p. 7705‐8.
362. Levine, T.P. and S. Munro, Targeting of Golgi‐specific pleckstrin homology domains involves
both PtdIns 4‐kinase‐dependent and ‐independent components. Current biology : CB, 2002.
12(9): p. 695‐704.
363. Godi, A., et al., FAPPs control Golgi‐to‐cell‐surface membrane traffic by binding to ARF and
PtdIns(4)P. Nature cell biology, 2004. 6(5): p. 393‐404.
364. Lenoir, M., et al., Structural basis of wedging the Golgi membrane by FAPP pleckstrin
homology domains. EMBO reports, 2010. 11(4): p. 279‐84.
365. Minogue, S., et al., Cloning of a human type II phosphatidylinositol 4‐kinase reveals a novel
lipid kinase family. The Journal of biological chemistry, 2001. 276(20): p. 16635‐40.
366. Waugh, M.G., et al., Localization of a highly active pool of type II phosphatidylinositol 4‐
kinase in a p97/valosin‐containing‐protein‐rich fraction of the endoplasmic reticulum. The
Biochemical journal, 2003. 373(Pt 1): p. 57‐63.
367. Guo, Y. and A.D. Linstedt, COPII‐Golgi protein interactions regulate COPII coat assembly and
Golgi size. J Cell Biol, 2006. 174(1): p. 53‐63.
368. Dowler, S., et al., Identification of pleckstrin‐homology‐domain‐containing proteins with
novel phosphoinositide‐binding specificities. Biochem J, 2000. 351(Pt 1): p. 19‐31.
369. Hughes, W.E., et al., SAC1 encodes a regulated lipid phosphoinositide phosphatase, defects in
which can be suppressed by the homologous Inp52p and Inp53p phosphatases. J Biol Chem,
2000. 275(2): p. 801‐8.
370. Godi, A., et al., ARF mediates recruitment of PtdIns‐4‐OH kinase‐beta and stimulates
synthesis of PtdIns(4,5)P2 on the Golgi complex. Nat Cell Biol, 1999. 1(5): p. 280‐7.
371. Audhya, A., M. Foti, and S.D. Emr, Distinct roles for the yeast phosphatidylinositol 4‐kinases,
Stt4p and Pik1p, in secretion, cell growth, and organelle membrane dynamics. Mol Biol Cell,
2000. 11(8): p. 2673‐89.
372. Shaywitz, D.A., et al., COPII subunit interactions in the assembly of the vesicle coat. J Biol
Chem, 1997. 272(41): p. 25413‐6.
373. Espenshade, P., et al., Yeast SEC16 gene encodes a multidomain vesicle coat protein that
interacts with Sec23p. J Cell Biol, 1995. 131(2): p. 311‐24.
374. Gimeno, R.E., P. Espenshade, and C.A. Kaiser, COPII coat subunit interactions: Sec24p and
Sec23p bind to adjacent regions of Sec16p. Mol Biol Cell, 1996. 7(11): p. 1815‐23.
375. Yorimitsu, T. and K. Sato, Insights into structural and regulatory roles of Sec16 in COPII
vesicle formation at ER exit sites. Molecular biology of the cell, 2012.
376. Connerly, P.L., et al., Sec16 is a determinant of transitional ER organization. Curr Biol, 2005.
15(16): p. 1439‐47.
377. Watson, P., et al., Sec16 defines endoplasmic reticulum exit sites and is required for secretory
cargo export in mammalian cells. Traffic, 2006. 7(12): p. 1678‐87.

78
378. Bhattacharyya, D. and B.S. Glick, Two mammalian Sec16 homologues have nonredundant
functions in endoplasmic reticulum (ER) export and transitional ER organization. Mol Biol
Cell, 2007. 18(3): p. 839‐49.
379. Misawa, H. and M. Yamaguchi, Molecular cloning and sequencing of the cDNA coding for a
novel regucalcin gene promoter region‐related protein in rat, mouse and human liver. Int J
Mol Med, 2001. 8(5): p. 513‐20.
380. Sawada, N. and M. Yamaguchi, Overexpression of RGPR‐p117 enhances regucalcin gene
promoter activity in cloned normal rat kidney proximal tubular epithelial cells: involvement of
TTGGC motif. J Cell Biochem, 2006. 99(2): p. 589‐97.
381. Sawada, N. and M. Yamaguchi, Overexpression of RGPR‐p117 enhances regucalcin gene
expression in cloned normal rat kidney proximal tubular epithelial cells. Int J Mol Med, 2005.
16(6): p. 1049‐55.
382. Iinuma, T., et al., Mammalian Sec16/p250 plays a role in membrane traffic from the
endoplasmic reticulum. J Biol Chem, 2007. 282(24): p. 17632‐9.
383. Hughes, H., et al., Organisation of human ER‐exit sites: requirements for the localisation of
Sec16 to transitional ER. J Cell Sci, 2009. 122(Pt 16): p. 2924‐34.
384. Ivan, V., et al., Drosophila Sec16 mediates the biogenesis of tER sites upstream of Sar1
through an arginine‐rich motif. Mol Biol Cell, 2008. 19(10): p. 4352‐65.
385. Whittle, J.R. and T.U. Schwartz, Structure of the Sec13‐Sec16 edge element, a template for
assembly of the COPII vesicle coat. J Cell Biol. 190(3): p. 347‐61.
386. Hughes, H. and D.J. Stephens, Sec16A defines the site for vesicle budding from the
endoplasmic reticulum on exit from mitosis. J Cell Sci, 2010. 123(Pt 23): p. 4032‐8.
387. Zacharogianni, M., et al., ERK7 is a negative regulator of protein secretion in response to
amino‐acid starvation by modulating Sec16 membrane association. EMBO J, 2011. 30(18): p.
3684‐700.
388. Witte, K., et al., TFG‐1 function in protein secretion and oncogenesis. Nature cell biology,
2011. 13(5): p. 550‐8.
389. Gimeno, R.E., P. Espenshade, and C.A. Kaiser, SED4 encodes a yeast endoplasmic reticulum
protein that binds Sec16p and participates in vesicle formation. J Cell Biol, 1995. 131(2): p.
325‐38.
390. Yamaguchi, M., Novel protein RGPR‐p117: its role as the regucalcin gene transcription factor.
Mol Cell Biochem, 2009. 327(1‐2): p. 53‐63.
391. Yamaguchi, M., S. Tomono, and T. Nakagawa, Overexpression of RGPR‐p117 suppresses
apoptotic cell death and its related gene expression in cloned normal rat kidney proximal
tubular epithelial NRK52E cells. Int J Mol Med, 2007. 20(4): p. 565‐71.
392. Yonekawa, S., et al., Sec16B is involved in the endoplasmic reticulum export of the
peroxisomal membrane biogenesis factor peroxin 16 (Pex16) in mammalian cells. Proc Natl
Acad Sci U S A, 2011. 108(31): p. 12746‐51.
393. Inoue, H., et al., Roles of SAM and DDHD domains in mammalian intracellular phospholipase
A1 KIAA0725p. Biochim Biophys Acta, 2012. 1823(4): p. 930‐9.
394. Mizoguchi, T., et al., Determination of functional regions of p125, a novel mammalian
Sec23p‐interacting protein. Biochem Biophys Res Commun, 2000. 279(1): p. 144‐9.
395. Aravind, L., The WWE domain: a common interaction module in protein ubiquitination and
ADP ribosylation. Trends in biochemical sciences, 2001. 26(5): p. 273‐5.
396. Zweifel, M.E., D.J. Leahy, and D. Barrick, Structure and Notch receptor binding of the tandem
WWE domain of Deltex. Structure, 2005. 13(11): p. 1599‐611.
397. Ponting, C.P., SAM: a novel motif in yeast sterile and Drosophila polyhomeotic proteins.
Protein science : a publication of the Protein Society, 1995. 4(9): p. 1928‐30.
398. Qiao, F. and J.U. Bowie, The many faces of SAM. Science's STKE : signal transduction
knowledge environment, 2005. 2005(286): p. re7.

79
399. Stapleton, D., et al., The crystal structure of an Eph receptor SAM domain reveals a
mechanism for modular dimerization. Nature structural biology, 1999. 6(1): p. 44‐9.
400. Thanos, C.D., K.E. Goodwill, and J.U. Bowie, Oligomeric structure of the human EphB2
receptor SAM domain. Science, 1999. 283(5403): p. 833‐6.
401. Kim, C.A., et al., Polymerization of the SAM domain of TEL in leukemogenesis and
transcriptional repression. The EMBO journal, 2001. 20(15): p. 4173‐82.
402. Harada, B.T., et al., Regulation of enzyme localization by polymerization: polymer formation
by the SAM domain of diacylglycerol kinase delta1. Structure, 2008. 16(3): p. 380‐7.
403. Nakajima, K., et al., A novel phospholipase A1 with sequence homology to a mammalian
Sec23p‐interacting protein, p125. J Biol Chem, 2002. 277(13): p. 11329‐35.
404. Yamashita, A., et al., Generation of lysophosphatidylinositol by DDHD domain containing 1
(DDHD1): Possible involvement of phospholipase D/phosphatidic acid in the activation of
DDHD1. Biochim Biophys Acta, 2010. 1801(7): p. 711‐20.
405. Sato, S., et al., Golgi‐localized KIAA0725p regulates membrane trafficking from the Golgi
apparatus to the plasma membrane in mammalian cells. FEBS Lett, 2010. 584(21): p. 4389‐
95.
406. Aviv, T., et al., The RNA‐binding SAM domain of Smaug defines a new family of post‐
transcriptional regulators. Nature structural biology, 2003. 10(8): p. 614‐21.
407. Bhunia, A., et al., NMR structural studies of the Ste11 SAM domain in the dodecyl
phosphocholine micelle. Proteins, 2009. 74(2): p. 328‐43.
408. Schuurs‐Hoeijmakers, J.H., et al., Mutations in DDHD2, encoding an intracellular
phospholipase A(1), cause a recessive form of complex hereditary spastic paraplegia. Am J
Hum Genet, 2012. 91(6): p. 1073‐81.
409. Willemsen, M.A., et al., Clinical, biochemical and molecular genetic characteristics of 19
patients with the Sjogren‐Larsson syndrome. Brain, 2001. 124(Pt 7): p. 1426‐37.
410. Shimoi, W., et al., p125 is localized in endoplasmic reticulum exit sites and involved in their
organization. J Biol Chem, 2005. 280(11): p. 10141‐8.
411. Ong, Y.S., et al., p125A exists as part of the mammalian Sec13/Sec31 COPII subcomplex to
facilitate ER‐Golgi transport. J Cell Biol, 2010. 190(3): p. 331‐45.
412. Arimitsu, N., et al., p125/Sec23‐interacting protein (Sec23ip) is required for spermiogenesis.
FEBS Lett, 2011. 585(14): p. 2171‐6.
413. Salama, N.R., T. Yeung, and R.W. Schekman, The Sec13p complex and reconstitution of
vesicle budding from the ER with purified cytosolic proteins. EMBO J, 1993. 12(11): p. 4073‐
82.
414. Salama, N.R., J.S. Chuang, and R.W. Schekman, Sec31 encodes an essential component of the
COPII coat required for transport vesicle budding from the endoplasmic reticulum. Mol Biol
Cell, 1997. 8(2): p. 205‐17.
415. Shugrue, C.A., et al., Identification of the putative mammalian orthologue of Sec31P, a
component of the COPII coat. J Cell Sci, 1999. 112 ( Pt 24): p. 4547‐56.
416. McGary, K.L., et al., Systematic discovery of nonobvious human disease models through
orthologous phenotypes. Proceedings of the National Academy of Sciences of the United
States of America, 2010. 107(14): p. 6544‐9.
417. Alam, I., et al., Differentially expressed genes strongly correlated with femur strength in rats.
Genomics, 2009. 94(4): p. 257‐62.
418. Sarmah, S., et al., Sec24D‐dependent transport of extracellular matrix proteins is required for
zebrafish skeletal morphogenesis. PLoS One, 2010. 5(4): p. e10367.
419. Rose, A.E., et al., Integrative genomics identifies molecular alterations that challenge the
linear model of melanoma progression. Cancer research, 2011. 71(7): p. 2561‐71.

Aim of the Project
80
This project aims to examine the following hypothesis:
p125A is a COPII specific accessory protein that regulates COPII assembly,
progression and activity through selective lipid binding
To address our hypothesis we took the following approaches:
1) Mapping and characterizing the specific lipid binding within the DDHD domain.
2) Examining and identifying the specific role of a Sterile α‐motif (SAM) assumed
responsible for p125A oligomerization, in particular in relation p125A lipid
recognition.
3) Examining the influence of wt and mutant forms of the SAM and DDHD regions on
lipid recognition in vitro.
4) Examining the influence of wt and mutant forms of the SAM and DDHD regions
for p125A targeting, stability and function at ERES in vivo.
5) Examining the role of p125A as a linker of the inner and outer COPII layers in
relation to the scaffolding activity of Sec16A.
6) Examining the function of the unstructured P‐Q‐rich N‐terminus of p125A.
7) Examining the ER and ERES targeting of mammalian Sec16A and Sec16B.
The results of the first five aims are presented in the manuscript entitled "Assembly of ER
exit sites is regulated by interactions of p125A with lipid signals", which has been submitted
to the Journal of Cell Biology. The results of the last two aims are presented in the chapter
“Investigations of p125A‐Sec31A associations and mammalian Sec16A and B membrane
binding”.

81
40040 characters
Assembly of ER exit sites is regulated by interactions of
p125A with lipid signals.
David Klinkenberg, Kimberly R. Long, Kuntala Shome, Simon C. Watkins and Meir Aridor &
& Correspondence
(Tel) 412‐624‐1970
e‐mail aridor@pitt.edu
Department of Cell Biology
University of Pittsburgh School of Medicine
3500 Terrace St. Pittsburgh PA 15261

Abstract
The inner Sar1‐Sec23/24 cargo‐sorting layer and the outer Sec13/31 cage layer of the
COPII coat mediate cargo sorting and vesicle biogenesis. mSec16A and p125A proteins
interact with both outer and inner coat layers to control coat activity yet the steps
directing functional assembly at ER exit sites (ERES) remain undefined. We hypothesize
82
that p125A utilizes lipid signals to control coat assembly. Within p125A, we defined a C‐
terminal DDHD domain found in phospholipases and PI transfer proteins that recognized
PA and phosphatidylinositol‐phosphates (PIP) in vitro and was targeted to PI4P‐rich
membranes in cells. A conserved central SAM domain promoted the assembly and
selective lipid recognition of the DDHD domain. A basic cluster and a hydrophobic
interface in the DDHD and SAM domains respectively were required for lipid recognition
and functional ERES assembly. The SAM‐DDHD lipid recognition module was utilized to
stabilize membrane binding and direct the spatial segregation of COPII from mSec16A,
nucleating the coat at ERES for ER exit.

Introduction
The COPII coat is composed of the small GTPase Sar1 and the cytosolic Sec23/24 and
Sec13/31 protein complexes (Antonny and Schekman, 2001). These cytosolic proteins
83
assemble on ER membranes to interact with and select cargo proteins destined for exit and
to deform membranes into buds and vesicles released from the ER. The coat inner layer,
Sar1‐Sec23/24, binds acidic lipids and presents multiple binding sites for short peptide
sequences, ER exit motifs, thus selecting cargo for incorporation into budded vesicles
(Aridor et al., 2001; Aridor et al., 1998; Kuehn et al., 1998; Miller et al., 2003). The outer
layer composed of the Sec13/31 complex forms an ancestral coat element 1 (ACE1) (Fath et
al., 2007) that is recruited on the inner layer and can polymerize to form an
icosidodecahedral cage (Stagg et al., 2008). A vesicle neck is constricted by the activity of
Sar1 leading to a GTPase dependent vesicle release (Bielli et al., 2005; Lee et al., 2005; Long
et al., 2010). This minimal set of COPII coat proteins recapitulates both cargo selection and
vesicle formation from ER membranes or synthetic liposomes (Antonny et al., 2001;
Matsuoka et al., 1998). However in vivo, COPII basic activities measured in minimal
reactions are controlled by interacting proteins that couple sorting and budding activities
with physiological biosynthetic demands (Zanetti et al., 2011).
The adaptation of ERES activities to changes in cargo load is mediated by the activities of
mSec16A and phosphoinositide 4‐kinase (PI4K) III (Farhan et al., 2008). Sec16p, an ACE1
containing protein, interacts with all COPII subunits and inhibits the GTPase activity of Sar1
by hindering proper linkage between the coat inner and outer layers (Yorimitsu and Sato,
2012) (Kung et al., 2011; Whittle and Schwartz, 2010). However, Sec16p substitutes for

84
acidic lipids in promoting COPII recruitment to synthetic liposomes and potentiates vesicle
formation on ER membranes (Supek et al., 2002).
PI4KIII generates phosphatidylinositol 4‐phosphate (PI4P) on ER membranes where the
dynamic generation of PI4P supports ERES assembly and ER export (Blumental‐Perry et al.,
2006; Farhan et al., 2008). The mechanisms by which these two activities, Sec16‐COPII
interactions and PI4P generation, control COPII budding remain to be defined.
p125A belongs to a family of PA preferring phospholipase A1 enzymes and was identified as
a Sec23 binding protein (Mizoguchi et al., 2000; Shimoi et al., 2005; Tani et al., 1999). p125A
associates with Sec31 in cytosol and is recruited to membranes with the Sec13/31 complex
where it binds Sec23, linking the two coat layers (Ong et al., 2010). Knockdown and over
expression studies demonstrate that p125A is required for ERES organization (Iinuma et al.,
2007; Shimoi et al., 2005) and cargo export from the ER (Ong et al., 2010). We now identify
a molecular cascade in which p125A functions as a multi‐domain adaptor that decodes lipid
signals such as PI4P. Lipid binding by p125A is utilized to spatially displace mSec16A while
directing COPII nucleation at ERES, promoting ER exit.

Results
85
p125A is recruited with COPII to PI4P enriched liposomes.
We hypothesized that COPII‐associated proteins may utilize selective PI4P recognition as a
mechanism to organize the coat at ERES for budding. To identify such proteins, we analyzed
Sar1‐induced recruitment of COPII from cytosol to control or PI4P containing liposomes.
Liposomes were incubated with cytosol in the presence of constitutively active (Sar1 H79G ,
termed Sar1‐GTP) or inactive (Sar1 T39N , Sar1‐GDP) Sar1 proteins. The recruitment of COPII to
membranes was analyzed by measuring the binding of cytosolic Sec23 to liposomes, isolated
by floatation in sucrose gradients. Effective COPII recruitment from cytosol to liposomes
was dependent on Sar1 activation and required PI4P (Fig. 1A). These results are in
agreement with previous studies showing that acidic lipids are required for the recruitment
of purified yeast COPII proteins to synthetic membranes (Matsuoka et al., 1998).
We analyzed the liposome binding reaction for the presence of the COPII associated protein
p125A that regulates ERES assembly and ER export (Ong et al., 2010; Shimoi et al., 2005).
p125A associates with Sec31 in cytosol and binds membrane‐recruited Sec23 using separate
segments on its N‐terminus (Ong et al., 2010). In agreement, immunodepletion of p125A
from cytosol did not affect the Sar1 dependent recruitment of COPII inner layer Sec23/24
from cytosol to ER membranes (Fig. 1B). Importantly, the p125A‐Sec31 complex was
recruited with Sec23 onto floated liposomes in Sar1 and PI4P dependent manner (Fig. 1C).
We thus hypothesized that p125A utilizes lipid recognition to regulate ERES assembly.

86
The DDHD and SAM domains cooperate to support lipid recognition in vitro and binding of
PI4P‐rich membranes in cells.
We hypothesized that p125A is a multi‐domain lipid‐regulated COPII adaptor. To test
possible roles for p125A in lipid recognition, we analyzed the lipid recognition properties of
selected p125A domains in isolation. The membrane binding characteristics of p125A reside
in the C‐terminus, which contains a DDHD domain and a sterile alpha motif (SAM). DDHD
domains are ~180 residues long (residues 779‐989 in p125A) and contain four conserved
residues (DDHD) that can form a putative metal binding site typically found in
phosphoesterase domains. DDHD domains are found in retinal degeneration B proteins, the
N‐terminal domain‐interacting receptor (Nir1‐3), where Nir2 functions as a PI‐transfer
protein (Litvak et al., 2005) and in the p125A‐containing PLA1 phospholipase protein family
(Sato et al., 2010; Shimoi et al., 2005; Yamashita et al., 2010).
In the context of many multidomain proteins, SAM domains are common protein‐protein
interaction motifs that modulate function through their ability to homo‐ or hetero‐
associate. Several forms of SAM domains have also been shown to polymerize into larger
functional structures (Qiao and Bowie, 2005). While commonly found in signaling and
nuclear proteins, SAM domains are also found in lipid modifying enzymes involved in
vesicular traffic (Nakatsu et al., 2010) (Nagaya et al., 2002a).
We analyzed the role of p125A SAM and DDHD domains in vivo and in vitro. In vivo, we
examined the cellular localization of selected EGFP tagged domains in transiently
transfected HeLa cells. In vitro, we analyzed the role of the domains in lipid recognition and
oligomerization using lipid‐blot overlays and sedimentation assays. As previously shown for
endogenous p125A (Shimoi et al., 2005), EGFP‐tagged p125A localized with COPII at ERES

(marked by the outer layer Sec31 subunit) that normally distribute either in the cell‐
periphery or clustered near the microtubule‐organizing center (MToC, Fig. 2A). At these
latter sites, ERES were adjacent but did not localize with the ER to Golgi intermediate
87
compartment (ERGIC53, Fig. 2A), cis (gpp130, Fig. 2A) or the trans‐Golgi network (TGN46,
not shown) compartments. At higher expression levels, EGFP‐p125A led to the enlargement
of COPII coated ERES membrane structures (Mizoguchi et al., 2000) (Figs. 7‐8). In contrast,
the isolated GFP tagged DDHD domain did not co‐localize with Sec31 but, strikingly, was
distributed in a cytosolic pool and also showed robust association with PI4P‐rich Golgi
membranes (Fig. 2B). Membrane bound EGFP‐DDHD decorated the rims of both cis (gpp130
not shown) and trans (TGN46, Fig. 2B) Golgi compartments adjacent to ERGIC, behaving like
typical PI4P reporters such as the PH domain of FAPP1 (Weixel et al., 2005).
We prepared a His6‐ tagged fragment of the C‐terminus encompassing the DDHD domain
(residues 701‐989) for analysis. Shorter fragments were difficult to produce because of low
yields indicating poor folding. In contrast with its targeting to PI4P‐rich membranes in cells,
when measured using lipid blot overlays the DDHD domain only displayed weak binding
with somewhat broad specificities toward acidic lipids including mono and poly
phosphorylated PIs, PA and PS (Fig. 3C). Given this ineffective lipid recognition, we produced
a larger fragment including both SAM and DDHD domains (residues 643‐989) for analysis.
Importantly, the combined domain exerted defined binding specificity to
monophosphorylated PIs including PI3P, PI5P and, in agreement with our cellular
observations, PI4P (Figs. 2B and 3D). The domain also recognized PA and PS. The results
suggest that inclusion of the SAM domain enhanced selective phospholipid recognition. It is
possible that the SAM domain binds selective lipids. Alternatively, it may assemble DDHD

domains to increase lipid‐binding avidity. To evaluate these possibilities, we generated a
88
GST‐tagged SAM domain (643‐701). Analysis using lipid‐blot overlay showed no lipid binding
(not shown). Furthermore, transiently expressed EGFP‐tagged SAM‐domain remained
cytosolic (Fig. 3F).
Structural studies have demonstrated that the homologous SAM domain of diacyl glycerol
kinase (DAGK) , a protein that regulates COPII assembly at ERES (Nagaya et al., 2002a),
dimerizes and generates oligomeric sheet structures that fall out of solution upon Zn 2+
binding. Zn 2+ ‐induced sheet formation is dependent on the ability of the domain to
dimerize, thus providing us with an easy test to monitor SAM oligomerization (Knight et al.,
2010). As observed with the SAM domain of DAGK, addition of Zn 2+ to the p125A‐SAM
domain led to robust polymerization of the protein, which quantitatively fell out of solution
under these conditions (Fig. 3H). Addition of Ca 2+ or Mn 2+ had minimal effects on SAM
solubility (not shown). The solubility of the control GST protein was also variably affected by
the addition of Zn 2+ . We thus cleaved the SAM domain from the GST. The isolated non‐
tagged SAM domain effectively precipitated in the presence of Zn 2+ whereas a dimer mutant
remained soluble (Fig. 3I, see below).
The lipid blot overlay analysis suggested that DDHD‐mediated lipid recognition is assisted by
p125A’s SAM domain, which may provide avidity‐based support for membrane binding. As
with EGFP‐DDHD, EGFP‐SAM‐DDHD localized to PI4P enriched Golgi and sometimes caused
Golgi disassembly (not shown) as observed with other PI4P binding domains (Weixel et al.,
2005). A GFP‐tagged fragment encompassing the linker region between the SAM and DDHD
domains (701‐779) remained cytosolic (not shown). Collectively, these results suggest a

model in which cooperative activities of assembled SAM and DDHD domains promote
89
selective lipid recognition and cellular membrane binding.
Segregation of ERES from ERGIC and Golgi at low temperatures reveals an exclusive
localization of p125A at ERES.
p125A may recognize monophosphorylated PIs including PI4P at ERES or alternatively on
ERGIC or Golgi membranes adjacent to COPII bud sites. To define the site of p125A‐
membrane binding, we analyzed the localization of p125A in relation to COPII, ERGIC and
Golgi markers in cells incubated at reduced temperatures that induce defined blocks in
anterograde and retrograde traffic between the ER and the Golgi. When cells were
incubated at 15C under conditions that arrest biosynthetic anterograde and retrograde
cargo traffic at ERGIC, ERGIC53 strongly accumulated in ERGIC compartments, with some
segregating into defined puncta as previously observed (Saraste and Svensson, 1991) (Fig.
4B). This transient distribution was rapidly reversed when returned to 37C and an
abundance of ERGIC53 containing tubular elements re‐clustered ERGIC at the MToC (not
shown). Golgi morphology remained largely unperturbed under these conditions (Fig. 4B).
Importantly, at 15C the number of ERES (marked by Sec31) was reduced while individual
sites were markedly enlarged and cytosolic COPII was effectively concentrated at these sites
(Fig. 4A).
Incubation of cells at 10C leads to the arrest of biosynthetic cargo at ERES (Mezzacasa and
Helenius, 2002). Under these conditions, ERES further coalesced at defined sites (Fig. 4A).
Importantly, at both 15C and 10C, p125A exclusively and dramatically partitioned with and
coated ERES (Fig. 4A). p125A coated ERES clearly segregated from both ERGIC and Golgi
compartments. The results suggest that at low temperatures, the typical organization of ER

90
exit complexes with ERES facing ERGIC containing VTCs was disrupted. Importantly, p125A
remained associated with enlarged COPII coated ERES to suggest that it resides exclusively
at ERES. Therefore, lipid recognition mediated by the co‐operative activity of SAM and
DDHD domains, which is required for p125A membrane binding, may occur at ERES.
COPII‐p125A containing ERES segregate from mSec16A at low temperatures.
Sec16p substitutes for the requirement of acidic lipids during COPII assembly on synthetic
membranes, while mSec16A and PI4KIII are both required during adjustment of ERES
assembly with cargo load (Farhan et al., 2008; Supek et al., 2002). We hypothesized that
lipid recognition by p125A may support COPII assembly at stages that precede or follow
Sec16 regulation. To explore this hypothesis, we localized Sec31, mSec16A and p125A using
the temperature blocks described above. We analyzed the localization of both endogenous
mSec16A (KIA00310, using specific antibody), as well as a GFP‐tagged mSec16A with relation
to the localization Sec31 and p125A with similar results (Fig. 5 and not shown). At 37C,
endogenous mSec16A and transiently expressed GFP‐mSec16A localized both at ERES and in
diffused cytosolic like localization. Importantly, imposing traffic blocks from the ER (10C) or
ERGIC (15C), led to robust collection of mSec16A at perinuclear sites adjacent but not
localizing with ERGIC or Golgi membranes (Fig. 5 and not shown). These results support a
dynamic distribution of mSec16A between cytosol and ER membrane, which is slowed down
by reduced temperatures.
Surprisingly, under these conditions, Sec31, Sec23 and mRFP‐p125A that localized to
enlarged peripheral ERES (Fig. 4A) all clearly segregated from mSec16A (Fig. 5 and not
shown). The robust segregation of Sec16 from COPII‐p125A coated ERES under conditions

that either block (10C) or slow (15C) cargo exit from the ER, suggest that p125A
participates in a late stage following the initial regulation of COPII by Sec16.
91
Charge and hydrophobic interactions are used by the SAM and DDHD domains to support
lipid recognition and assembly.
Our in vivo (Fig. 2B, 3B and F) and in vitro analysis (Fig. 3D) suggests that lipid recognition
resides within the DDHD domain and is assisted by SAM domain mediated assembly (Fig. 3F
and D). To test this hypothesis, we generated mutations in these domains and examined
their functionality. Within the DDHD domain we focused on a group of basic residues (851‐
KGRKR‐855) replacing those with glutamic acid (851‐EGEEE‐855, termed PI‐X). When tested
in lipid blot overlay assays, the His6‐tagged SAM‐DDHD PI‐X domain showed no lipid
recognition (Fig. 3E) and transiently expressed EGFP‐tagged DDHD PI‐X lost its Golgi
localization presenting a diffuse cytoplasmic distribution (Fig. 3B). This localization was also
observed when EGFP‐SAM‐DDHD PI‐X was analyzed (not shown). The results suggest that the
DDHD domain is required for lipid recognition.
We further hypothesized that the SAM domain, which does not display lipid recognition or
cellular targeting in isolation (Fig. 3F and not shown), supports protein assembly. To test this
hypothesis, we used structural information available for the DAGK ‐SAM domain to guide
mutagenesis aimed at abolishing protein assembly (Knight et al., 2010; Qiao and Bowie,
2005). In the DAGK ‐SAM domain, hydrophobic interactions between valine and leucine
residues mediate dimerization required for oligomerization of the domain (Fig. 3A and G),
whereas introduction of a charged residue at this position prevented both dimerization and
Zn 2+ ‐ induced high order oligomerization. The hydrophobic dimer interface is fully
conserved in p125A, thus we could generate a single residue replacement in p125A SAM‐

92
domain (L690E) for analysis. Unlike the wild type protein, GST‐SAM (L690E) did not precipitate
in response to Zn 2+ (Fig. 3H and I), thus in common with DAGK δ, this single point mutation
abolished dimerization and therefore high order assembly of the domain.
Similarly, while the untagged SAM domain robustly precipitated with Zn 2+ addition,
SAM (L690E) remained completely soluble. The results suggest that the hydrophobic assembly
interface within the SAM domain of p125A is functional.
Assembly controlled lipid‐recognition is required to regulate COPII organization at ERES.
We hypothesized that p125A utilizes lipid recognition to regulate COPII assembly at ER exit
sites. Tagged p125A in which specific residues required for SAM‐domain assembly and
DDHD supported lipid recognition (Fig. 3) were mutated (p125A PI‐X , p125A L690E and SAM and
DDHD double mutant p125A PI‐X, L690E ) were generated to test this hypothesis. Individual
mutations in the SAM (L690E) or DDHD domain (PI‐X) may not be sufficient to generate a
dominant phenotype because PI‐X mutants may assemble with the endogenous protein
while L690E mutants may retain lipid recognition (Figs. 2B and 3B). In both cases, p125A is
expected to maintain its interactions with both layers of COPII as these are mediated by the
unperturbed N‐terminus (Ong et al., 2010).
WT EGFP‐p125A localizes to ERES (Fig. 6). EGFP‐p125A PI‐X was also targeted to ERES,
however a diffused cytosolic component was quite evident. Over expression did not lead for
the most part to clustering as observed with the wild type protein (not shown). The
dimerization interface mutant EGFP‐p125A L690E (Fig. 3H and I) exhibited diffuse labeling and
association with ERES similar to the PI‐X mutant. However, overexpression of EGFP‐
p125A L690E led to augmented diffused cytosolic distribution with occasional targeting to both

93
ERES and the perinuclear Golgi region (Fig. 6). Importantly, in marked contrast to WT p125A,
the double mutant (p125A PI‐X, L690E ), in which the presumed cooperative lipid‐binding module
is disabled, showed no ERES localization and exhibited diffused cytosolic distribution, which
was maintained at very high expression levels (Figs. 6 and 8A). These results suggest that
selective lipid recognition mediated by SAM‐DDHD module is required for p125A‐membrane
binding at ERES.
We thus tested whether selective membrane binding by p125A regulates COPII assembly at
ERES. Indeed, p125A PI‐X, L690E became a trans‐dominant negative inhibitor of ERES assembly.
Sec31 lost ERES localization and remained diffusely localized in the cytoplasm (Fig. 6). These
results suggest that cooperative lipid recognition derived from SAM‐DDHD activities is
required to regulate COPII assembly at ERES. The DDHD domain recognized PI4P rich
membranes in cells and on lipid blot overlays. Similar to p125A PI‐X, L690E , deletion of the
DDHD domain (778‐989) in the background of SAM domain inactivation (using the L690E
mutation, p125A L690E, ΔDDHD ) led to cytosolic dispersion of the protein and inhibited ERES
assembly as analyzed by Sec31 staining (Fig.7C).
We used this truncation to further test the role of p125A SAM‐DDHD as a selective lipid
recognition module directing ERES assembly, by artificially replacing the domain with a bona
fide PI4P binding domain, the Pleckstrin homology (PH) domain of Fapp1 (Blumental‐Perry
et al., 2006). Fapp1‐PH confers selective recognition of PI4P and as observed with the
isolated DDHD domain, is targeted in isolation to PI4P‐rich Golgi membranes (Weixel et al.,
2005). When expressed in cells, the chimera (p125A L690E, ΔDDHD, +Fapp1‐PH ) displayed both
cytosolic and punctate configuration with some preferential targeting to the Golgi (Fig. 6).
Importantly, in contrast to expression of p125A L690E, PI‐X or p125A L690E, ΔDDHD , which disrupted

ERES assembly, the chimera maintained the assembly of endogenous Sec31 at both
peripheral ERES and perinuclear Golgi adjacent sites. The ability to artificially replace the
94
SAM‐DDHD lipid recognition module with a PI4P binding domain and restore ERES assembly,
supports the role of the selective lipid recognition and in particular the role of PI4P in p125A
mediated assembly of ERES.
Lipid recognition controls p125A residency at ERES
We further hypothesized that the increase in the cytosolic population seen with p125A PI‐X
and p125A L690E as well as the dispersed nature of the double mutant and Sec31 (Fig. 6) is
derived from reduced association of p125A mutants with ERES membranes. To test this
hypothesis, we analyzed the dynamics of p125A proteins and COPII at ERES using
fluorescence recovery after photobleaching (FRAP).
HeLa cells were transiently transfected with constructs expressing YFP‐Sec23, EGFP‐p125A
(Fig. S1) or mRFP‐p125A (not shown). An average of 27‐35 measured events for each time‐
point collected in three independent experiments is shown. COPII marked by both YFP‐
Sec23 (Fig. S1B) or CFP‐Sec31 (not shown) exhibited fast recovery kinetics as previously
reported (Forster et al., 2006) whereas EGFP‐p125A (Fig. S1C) or mRFP‐p125A (not shown)
both exhibited similarly or slightly slower dynamics. Importantly, in agreement with
morphological analysis (Fig. 6), a faster recovery was recorded for EGFP‐p125A L690E (Fig.
S1D), and EGFP‐p125A PI‐X (Fig. S1E) providing an explanation for the observed diffuse
cytosolic population of these mutants (Fig. 6). The data fitted well with a single exponential
showing an averaged T½ for YFP‐Sec23 of 3.14 Sec and for EGFP‐p125A, T½ of 3.36 Sec. In
contrast, EGFP‐p125A L690E (2.98 Sec) and EGFP‐p125A PI‐X (2.56 Sec) exhibited faster kinetics.

EGFP‐p125A L690E, PI‐X did not assemble at defined sites and hence could not be measured.
However, limited association with ERES was observed when EGFP‐p125A L690E, PI‐X was
95
analyzed at reduced temperatures (not shown) suggesting that enhanced turnover at ERES
prevented stable localization. These results suggest that selective lipid binding through the
SAM‐DDHD module controls p125A association with ERES membranes.
p125A functions at a late stage in ERES nucleation.
p125A‐Sec23‐Sec31 coated ERES segregated from mSec16 during temperature induced
traffic blocks at ERES or ERGIC (Figs. 4‐5). We hypothesized that p125A actively displaces
mSec16A from ERES, suggesting that p125A over‐expression may facilitate such
displacement. Over‐expression of mRFP‐p125A led to a marked uniform enlargement of
ERES as previously demonstrated (Figs. 7A, 8A). The large sites marked by ER membranes
containing the cargo protein Venus‐VSV‐G (Fig. 8C), collected both layers of COPII as
analyzed by the co‐localization of endogenous Sec31 or co‐expressed YFP‐Sec23 (Fig. 7B and
not shown).
We thus analyzed if GFP‐mSec16A is displaced from these sites. When expressed at low
levels, mRFP‐125A largely co‐localized with mSec16A, Sec31 and Sec23 at ERES (Fig. 5). In
contrast, when over‐expressed, p125A induced large ERES that were clearly lacking GFP‐
mSec16A (compare YFP‐Sec23‐mRFP‐p125A to GFP‐mSec16A‐mRFP‐p125A, Fig. 7A‐B).
Occasionally, mSec16A was found adjacent to these enlarged and somewhat rounded sites
that varied from 500‐1500 nm in size as shown by super resolution SIM microscopy (Fig. 8B).
Sec16p negates the requirements for acidic lipids in COPII binding to liposomes (Supek et al.,
2002). We hypothesized that p125A might similarly utilize PI4P binding to direct mSec16A

displacement thus stabilizing COPII‐membrane binding. We therefore analyzed if over‐
96
expressed p125A proteins that are deficient in lipid recognition are also defective in Sec16
displacement.
Two proteins were analyzed in which the lipid binding module was inactivated or deleted;
mRFP‐p125A L690E, PI‐X and p125A L690E, ΔDDHD . In marked contrast to mRFP‐p125A, mRFP‐
p125A L690E, PI‐X remained dispersed even at very high expression levels (Fig. 8A). Over‐
expressed p125A L690E, ΔDDHD also remained largely cytosolic but enlarged sites were now
occasionally evident (Figs. 7C‐D, 8A‐B). Importantly, these sites effectively collected GFP‐
mSec16A (Fig. 7D). Analysis using SIM microscopy demonstrated that p125A L690E, ΔDDHD ‐
induced sites were now effectively engulfed within GFP‐mSec16A (Fig. 8B, Movies S1‐2).
Thus mSec16A is displaced by p125A in a manner that is regulated by lipid binding.
The unusual morphology observed with p125A overexpression suggested that the coated
sites might become inhibitory to biosynthetic traffic. However, although the temperature
synchronized traffic of tsVSV‐G from the ER to the Golgi was inhibited by expression of both
WT or p125A mutants to a variable degree (most likely due to variable expression levels
between experiments, not shown), traffic was not blocked as analyzed morphologically and
by following the acquisition of resistance to endoglycosidase H digestion on tsVSV‐G (Fig. 8C
insert).
High‐resolution microscopy identified uncoated VSV‐G containing vesicular structures
budding off these enlarged sites (Fig. 8C, Movies S3‐5). The results suggest that ERES were
completely coated during p125A over expression yet allowed for the typical formation of
multiple cargo containing buds. The lack of coat on these buds is in agreement with previous

97
studies showing that the sites contained p115 (Mizoguchi et al., 2000) and thus progressed
beyond Sar1‐GTP hydrolysis and initial uncoating during vesicle release.
Functional contribution of the SAM‐DDHD membrane‐binding module.
Depletion of p125A using RNAi leads to disruption of ERES (Shimoi et al., 2005) and causes
kinetic inhibition of membrane and soluble cargo secretion from the ER leading to the
disruption of Golgi morphology (Ong et al., 2010). To examine the contribution of the SAM‐
DDHD lipid‐binding module to p125A activity, we replaced endogenous p125A with EGFP‐
p125A L690E, PI‐X . We analyzed Golgi morphology as a robust reporter for overall steady state
traffic activities. Golgi morphology was analyzed using gpp130 (Fig. 9) or GalNAcT2‐GFP
localization (Fig. S2). Morphology was heterogeneous with four distinct phenotypes (Fig.
9A): 1. Intact Golgi localized to the perinuclear region. 2. Loosely packed or dispersed Golgi
located in the perinuclear or around the nuclei. 3. Completely shattered (vesiculated) Golgi
dispersed throughout the cell. 4. Cells missing detectable Golgi compartment perhaps
reporting on mitotic cell population. Effective depletion of endogenous p125A was achieved
as previously reported (Ong et al., 2010)(Fig. 9B).
Depletion of p125A led to dramatic reduction in the intact Golgi population and a
concomitant increase in shattered morphology when compared to control RNAi treated cells
(Fig. 9C‐D). The effect was specific as expression of an RNAi resistant form of EGFP‐p125A
(Fig. 9B‐D) reversed these effects, namely restoring intact Golgi populations and eliminating
L690E, PI‐X
the shattered Golgi morphology (Fig. 9C‐D). In contrast, expression of EGFP‐p125A
was ineffective in correcting Golgi morphology, leading only to a partial restoration of intact
morphology and reduction in shattered Golgi compartments (Fig. 9C‐D). Overall these
results suggest that defects in the activity of the SAM‐DDHD lipid‐binding module of p125A,

98
which led to robust morphological defects in ERES assembly (Fig. 6) reduced association of
p125A with ERES (Fig. S1) and inhibition of Sec16 segregation from ERES (Figs. 5, 7‐8) also
translated into functional defects in steady state traffic activities required to maintain Golgi
morphology.
Discussion
While COPII core subunits are sufficient in mediating vesicle biogenesis, COPII interacting
proteins control budding activities at ERES. We defined a cascade in which p125A, a protein
that links both COPII layers, utilizes a lipid recognition module (Figs. 1‐3) to stabilize
membrane‐binding (Fig. S1) while promoting the spatial segregation of COPII from mSec16A
(Figs. 4‐5, 7‐8), leading to functional ERES assembly (Figs. 6, 9). Thus, regulatory activities
such as lipid signaling (Blumental‐Perry et al., 2006; Farhan et al., 2008; Nagaya et al.,
2002b; Pathre et al., 2003) control the progression of a pre‐budding cascade that directs
COPII nucleation and activity at ERES (Fig. 10).
The SAM‐DDHD lipid‐binding module.
Previous studies suggested that the C‐terminus of p125A, which contains SAM and DDHD
domains, is involved in membrane binding. A defined function of SAM domains is protein
oligomerization (Qiao and Bowie, 2005), used to increase avidity between assembled
complexes and substrates. Our findings suggest the basic assembly activity of SAM is
functional in p125A. First, we showed that in common with its close homolog, the SAM
domain of DAGK (Knight et al., 2010), the p125A‐SAM domain oligomerized when bound to
Zn 2+ . Second, we demonstrated that the basic assembly interface is conserved and
introduction of a single point mutation within the site abolished oligomerization (Fig. 3).

Third, we demonstrated that the SAM assembly interface within p125A is required to
99
support ERES organization and activity (Figs. 6‐9 and S1).
Because p125A‐SAM lacked membrane targeting in vivo or lipid‐binding activities in vitro,
we favor a model in which this domain enhances the avidity for lipid recognition by the
DDHD domain as suggested by lipid blot overlay analysis (Fig. 3). This activity may be shared
in other lipid sensing and processing enzymes that function in vesicular transport. In DAGK,
a SAM‐PH module is required for inhibition of ERES assembly although lipid‐binding
specificities of this PH domain are undefined (Nagaya et al., 2002b). The SAM domain of the
inositol 5‐phosphatase Ship2 that regulates clathrin mediated endocytosis may be similarly
required for membrane recognition (Nakatsu et al., 2010). The Ship2‐SAM domain further
supports hetero dimerization with the SAM domain of the PI3‐kinase effector Arap3 that
contains PH domains and serves as a GTPase activating protein (GAP) for both Arf and Rho
G‐proteins (Raaijmakers et al., 2007). By analogy, heterodimeric interactions between SAM‐
DAGK and SAM‐p125A may function to form lipid‐recognition and processing hubs at ERES.
Lipid recognition most likely resides in the p125A‐DDHD domain. When expressed in cells in
isolation, EGFP‐DDHD was targeted to PI4P‐rich Golgi membranes (Fig. 2). Mutations in the
domain (DDHD PI‐X ), which prevented Golgi binding in cells (and further abolished the
targeting of EGFP‐SAM‐DDHD PI‐X domain, not shown), also abolished lipid recognition by the
SAM‐DDHD module in vitro (Fig. 3). These mutations probably did not destabilize DDHD PI‐X
because it did not aggregate in cells and was produced in bacteria at yields higher than its
WT version. Moreover, deletion of the DDHD domain (p125A L690E, DDHD ) abolished
membrane binding and dispersed ERES similarly to proteins carrying the DDHD PI‐X mutations
(p125A L690E, PI‐X , Figs. 6‐7). Future structural studies are needed to define the contributions of

100
the basic PI‐X cluster in lipid recognition. Together, the SAM‐DDHD module provides a lipid
recognition unit for p125A that regulates COPII assembly (Fig. 6).
In vitro analysis suggests that the SAM‐DDHD module has somewhat broad lipid binding
specificity that includes monophosphorylated PIs and PA (Fig. 3). However several lines of
evidence suggest that PI4P might be the primary target for this module. 1. Full‐length p125A
shows specificity for PIP recognition (Inoue et al., 2012; Shimoi et al., 2005). 2. p125B, a
family member of p125A that localizes to the Golgi, recognizes PI4P using its SAM and DDHD
domains although individual contributions of these domains were not defined (Inoue et al.,
2012). p125B‐SAM‐DDHD can replace the p125A lipid recognition module and the resulting
chimera localizes at ERES (Shimoi et al., 2005). 3. GFP‐DDHD domain is targeted to PI4P rich
Golgi membranes (Figs 2‐3). 4. p125A with disabled SAM dimer interface (L690E mutant)
and mutated DDHD domain (PI‐X) loses membrane targeting leading to ERES disassembly as
observed when p125A L690E, ΔDDHD was examined (Figs. 6‐7). Targeting as well as ERES
organizing activity was partially restored by simple replacement of the DDHD domain with a
well‐characterized PI4P‐binding domain (Fapp1‐PH, Fig. 6). 5. Other DDHD domain
containing proteins including p125B (Nakajima et al., 2002; Yamashita et al., 2010) and Nir‐2
(Litvak et al., 2005) are localized to PI4P‐rich Golgi membranes. Similarly, p125A may utilize
PI4P to localize and regulate COPII activities at ERES.
Role of p125A in ERES regulation.
A dependency of yeast COPII assembly on acidic lipids and in particular PI4P was originally
deduced from analysis using synthetic membranes. This dependency is abrogated by the
inclusion of Sec16p in such reactions providing the first link between Sec16p and lipid
signals (Supek et al., 2002). However, depletion of the yeast major PI4‐kinases PIK1 and Stt4

101
fails to affect ER to Golgi traffic (Audhya et al., 2000). Subsequent studies have shown that
sequestration of Golgi PI4P in vitro or prolonged PIK1 inactivation in vivo, do inhibit ER to
Golgi traffic (Lorente‐Rodriguez and Barlowe, 2011) yet inhibition is exerted on fusion of
COPII vesicles with Golgi membranes. Unlike yeast, mammalian COPII vesicles do not fuse
with the Golgi but rather fuse homotypically in the vicinity of bud sites, suggesting that PI4P
formation is initiated at these sites. Indeed in mammals, PI4P is utilized to regulate ERES
assembly and ER export (Blumental‐Perry et al., 2006; Farhan et al., 2008).
We suggest that Sec16 and PI4P‐p125A interactions represent sequential steps in the COPII
budding cascade (Fig. 10). Several lines of evidence support this model. 1. mSec16A is
required to nucleate new ERES, and both mSec16A and the ER‐localized PI 4‐kinase type III
are required to maintain these sites (Farhan et al., 2008). 2. Sec16p‐COPII interactions
hinder proper linkage between COPII layers thus inhibiting Sec31 stimulated GAP activity
(Kung et al., 2011; Supek et al., 2002; Yorimitsu and Sato, 2012). Proper linkage is required
for the completion of vesicle budding whereas Sec16p is dispensable in such reactions
(Fromme et al., 2007). 3. mSec16A is slightly removed from bud sites at steady state
(Hughes et al., 2009), whereas at low temperatures that block ER exit at late stages, it is
displaced from arrested ERES (Fig. 5). 4. mSec16A displacement is further enhanced by over
expression of p125A (Fig. 7‐8).
Our model (Fig. 10) suggests that p125A, which interacts with both COPII layers, may
provide a mechanism to facilitate the progression of COPII budding, promoting Sec16
dissociation while linking both COPII layers. p125A utilizes its SAM‐DDHD lipid binding
module in these reactions (Fig. 7‐8) thus employing lipid signals to facilitate the progression
of the ERES assembly cascade.

102
Sec16 and p125A interact with the Sec31 ‐solenoid containing c‐terminal domain (Ong et
al., 2010; Shaywitz et al., 1997). How p125A‐membrane binding regulates these interactions
to promote the segregation of mSec16A from ERES remains to be defined. In yeast, acidic
lipids support inner layer binding to membranes non selectively thus negating the need for
Sec16p (Matsuoka et al., 1998) while in higher eukaryotes, selective lipid recognition by
p125A provides regulation of COPII assembly (Fig. 10).
The physiological role of p125A remains to be determined. Acute depletion or over‐
expression of p125A leads to general traffic defects (Figs. 8‐9). p125A depletion interferes
with neural crest cells migration in Xenopus and was predicted to be a causative gene in the
development of Waardenburg syndrome (McGary et al., 2010). However p125A‐KO mice are
relatively unaffected (Arimitsu et al., 2011), mainly presenting defects in spermiogenesis.
Sec16 may provide sufficient support for COPII activities in these animals as observed in
vitro by the functionality of COPII on liposomes that contain Sec16p yet lack acidic lipids
(Supek et al., 2002). The identified steps in ERES assembly are likely subjected to
physiological regulation, which remains to be defined.

Materials and Methods.
103
HeLa cells were all maintained at sub‐confluence in Dulbecco's Modified Eagle's Media (DMEM) (HyClone Fisher‐Scientific)
supplemented with up to 10 % Fetal Bovine Serum (FBS)(Serum Source International, Inc.) and 5 % Penicillin‐Streptomycine
(Cellgro) under standard incubation environment (37°C, 5 % CO2). Antibodies used in the study include rabbit anti p125A
antibody against the N‐terminus (300‐592) of p125A (MSTP053, Bethyl Lab), rabbit anti p125A antibody against the C‐
terminus (732‐752) of p125A (AP114511b, Abgent), mouse monoclonal anti p125A and rabbit anti KIAA03100 (Sec16L) kindly
provided by Dr. Katsuko Tani (School of Life Science, Tokyo University of Pharmacy and Life Science, Hachioji, Tokyo, Japan).
Mouse monoclonal against Sec31A (612350) was from BD Transduction Laboratories. All Golgi specific antibodies were kindly
provided by Dr. Adam Linstedt (Department of Biological Sciences, Carnegie Mellon University, Pittsburgh, PA, USA). Rabbit
polyclonal against GFP was from Polysciences Inc (Cat # 24240). Mouse monoclonal against ERGIC53 (G1/93, ALX‐804‐602)
was from Enzo Life Science and Mouse monoclonal against Flag (M2, F1804) was from Sigma‐Aldrich. Mouse monoclonal
against β‐Actin (ab6276, Abcam), Mouse monoclonal against GFP (332600, Invitrogen). All Alexa‐conjugated goat anti mouse
or rabbit antibodies were from Invitrogen. HRP‐conjugated rabbit against GST (ab3416‐250, Abcam). HRP‐conjugated Mouse
against His (040905270001, Roche). All HRP‐coupled secondary antibodies were from Pierce.
p125A was excised from pFlag‐CMW‐6c‐p125A (kindly provided by Dr. Katsuko Tani, School of Life Science, Tokyo
University of Pharmacy and Life Science, Hachioji, Tokyo, Japan) using unique Hind III and Sma I restriction sites and ligated
into the same sites of pEGFP‐C1H, a modified pEGFP‐C1 derivative where a Hind III site in the MCS had been added in‐
frame. EGFP‐mSec16L was kindly provided by Dr. Vivek Malhotra GRC, Spain.
Selected p125A domains were all PCR amplified while introducing a Hind III (5') or a stop codon followed by a Sma I site (3')
p125 Hind III aa:643 F: 5'‐CGTATGACCTTGTTAAGCTTAATAAAGAAGTCCTAACTTTGC‐3'
p125 Hind III aa:779 F: 5'‐GGACAGGTTTCTGTTGCTTACAAGCTTTTAGATTTTGAACCAGAGATATTCTTTGC‐3'
p125 Hind III aa: 701 F: 5'‐CCCAGAAAGAAGATAGCTAACAAGCTTGAACATAAAGCAGCC‐3'
p125 aa:704 Stop Sma I R: 5'‐CCTTCTTTTCTGACGCTGCTTTTTTCCCGGGTTATGCTTTATGTTCTACAAAGTTAGC‐3'
p125 (L779Stop) Sma I R: 5'‐ GCAAAGAATATCTCTGGTTCCCCGGGTTATGAGTTGTAAGCAACAGAAACC‐3'
p125 aa:990 Stop Sma I R: 5'‐GGGGCTGTTCTGGACTACCCGGGAATCATCGATAAATTTCTTTAAGTAGTAACAGAGC‐3'

104
The p125A L690E, PI‐X (850KGRKREGEEE854) and DsRNAi resistance mutations were introduced by 2 Step PCR
mutagenesis.
p125 L690E (SAMX) F: 5'‐CCTGAAGGAAATGGGGATACCCGAAGGACCCAGAAAGAAGATAGC‐3'
p125 L690E (SAMX) R: 5'‐GCTATCTTCTTTCTGGGTCCTTCGGGTATCCCCATTTCCTTCAGG‐3'
p125 850(KGRKR/EGEEE)854 F:
5'‐GGACCTAAAAGCTGTTCTCATTCCACATCACGAAGGCGAAGAAGAACTTCATTTAGAATTGAAAGAGAGTCTCTCTCG‐3'
p125 850(KGRKR/EGEEE)854 R:
5'‐CGAGAGAGACTCTCTTTCAATTCTAAATGAAGTTCTTCTTCGCCTTCGTGATGTGGAATGAGAACAGCTTTTAGGTCC‐3'
p125 R III F: 5'‐GGAGATGCCTCAAGTTGACCACCTAGTCTTCGTGGTGCATGGCATTGGACCTGTGTGTG‐3'
p125 R III R: 5'‐ CACACACAGGTCCAATGCCATGCACCACGAAGACTAGGTGGTCAACTTGAGGCATCTCC‐3'
The p125A‐Fapp1‐PH chimera was constructed by excising the DDHD domain of pEGFP‐p125A using flanking Pvu I
restriction sites that were introduced by PCR. The Fapp1‐PH domain was amplified from pGEX‐4T‐1‐Fapp1‐PH introducing
flanking Pvu I sites and inserted into the introduced Pvu I sites:
Δp125 (DDHD) Pvu I Ins F: 5'‐GAAATTCGATCGACAATGAACATTAGTCCAGAACAGC‐3'
Δp125 (DDHD) Pvu I Ins R: 5'‐GGTTCAAACGATCGTGAGTTGTAAGCAACAGAAACC‐3'
Fapp1‐PH Pvu I F: 5'‐GGTTCCGCGTGGATCCCCGCGATCGATGGAGGGGGTGTTG‐3'
Fapp1‐PH Pvu I R: 5'‐CACGATGCGGCCGCTCGCGATCGCTTAGTCCTTGTATCAGTCAAAC‐3'
pmRFP‐C1H: pmRFP‐C1H vector was created by replacing EGFP ORF in pEGFP‐C1H with the mRFP ORF from pmRFPSec61β,
kindly provided by Dr. Tom A. Rapoport (Harvard Medical School, Boston, MA), using the flanking Nhe I and Xho I sites in
both vectors. p125A constructs were subcloned into pmRFP‐C1H using the Hind III and Sma I restriction sites. The pmRFP‐
p125 L690E‐Fapp1‐PH was generated by introducing the L690E mutation into the pmRFP‐p125A‐Fapp1‐PH by 2 step PCR
mutagenesis. The pmRFP‐p125∆DDHD‐L690E expression construct was generated by excision of the Fapp1‐PH through
digestion of the pmRFP‐p125 L690E‐Fapp1‐PH with PvuI.

105
The SAM domain was amplified from pEGFP‐p125A using (5’)Hind III‐ ‐ (3’) Stop‐Sma I containing primers as described
above and introduced into pGEX‐4T‐1H, a derivative of pGEX‐4T‐1 (GE Healthcare Life Science) where a unique Hind III had
been added in‐frame with GST. p125A SAM‐DDHD or DDHD (643‐989 or 701‐989) were PCR amplified out of pEGFP‐p125A
(WT or PI‐X) while introducing a 5' Nde I site and Stop codon followed by a Hind III site at the 3'. The fragments were
cloned into pET28a(+) bacterial expression vector (Novagen) to generate His6‐tagged domains.
p125 Nde I aa: 643 F: 5'‐CGTATGACCTTGTTCATATGAATAAAGAAGTCCTAACTTTGC‐3'
p125 Nde I aa: 701 F: 5'‐ CGTCAGAAAAGAAGGCAGTGGCGCATATGGAACATAAAGCAGCC‐3'
p125 Nde I aa: 779 F: 5'‐GGACAGGTTTCTGTTGCTTACCATATGTTAGATTTTGAACCAGAGATATTCTTTGC‐3'
p125 (L779Stop) Hind III R: 5'‐ GCAAAGAATATCTCTGGTTCAAGCTTTTATGAGTTGTAAGCAACAGAAACC‐3'
p125 aa: 990 Stop Hind III R: 5'‐GGGGCTGTTCTGGACTCTAAAGCTTTCATCGATAAATTTCTTTAAGTAGTAACAGAGC‐3'
All Clones were verified by sequencing (Genewiz).
Transfection was carried out using Effectene (Qiagen) or Lipofectamine 2000 (Life Sciences) transfection reagents
according to provided protocol, with optimized DNA concentrations. ER microsomes were prepared from NRK cells as
previously described (Rowe et al., 1996). His6 tagged Sar1 H79G and T39N proteins were purified as previously described
(Aridor et al., 1995; Rowe and Balch, 1995). Rat liver cytosol was prepared as described (Aridor et al., 1995). His6 tagged
DDHD, SAM‐DDHD and SAM‐DDHD PI‐X were purified on Ni‐NTA‐Agarose (Qiagen) using a Sarcosyl extraction protocol
(Frangioni and Neel, 1993). Briefly, protein expression in transformed BL 21DE3 (Invitrogen) was induced with IPTG (0.1
mM) for 4 hr at 37C and cells were collected and lysed as previously described (Aridor et al., 1995). 10 % N‐Lauroyl
Sarcosine (MP Biomedicals) was added to cell lysates which were further homogenized by sonication. Cell debris were
removed by centrifugation and supernatants were supplemented with 2 % n‐Octyl‐β‐D‐glucopyranoside (OG) (Gold
Biotechnology USA). Solubilized proteins were loaded on Ni‐NTA‐Agarose (Qiagen). Protein bound beads were washed
three times with buffer containing 50 mM Tris‐HCl (pH=8.0), 100 mM NaCl, 1 mM EDTA, 1 mM PMSF and 2 % OG, followed
by three washes with HNE buffer (50 mM HEPES (pH=7.4), 300 mM NaCl, 1 mM MgCl2, 0.5 mM EGTA, 2 % OG) and three
washes with HNE buffer supplemented with 25 mM Imidazole (pH=7.4). Bound proteins were eluted with HNE buffer
containing 500 mM Imidazole . GST‐ tagged SAM and SAM L690E proteins were purified on GS‐Sepharose 4B (GE Healthcare
Life Science) and thrombin cleaved using the standard bulk GST purification protocol.

p125A knockdown‐replacement analysis.
106
HeLa cells were plated into 6‐Well Plates at a density of 2 x 10 5 u / well and incubated overnight. Subsequently DsRNAi (200
nM , IDT) were transfected using Oligofectamine (Invitrogen) according to provided protocols. The transfection procedure
was repeated after 24 hr. and the cells were incubated for additional 12 hr. Each treated well was subsequently expanded
into 4 wells and left to incubate for 12 hr. before transfection with EGFP‐p125A resistant clones. EGFP‐p125A resistant clones
were expressed for 12‐14 hr. and were processed for IF or WB. The following p125A DsRNAi were obtained from IDT
according to (Shimoi et al., 2005):
5'‐rArArGrUrUrGrArCrCrArUrUrUrGrGrUrGrUrUrUrGrUrGrdGdT‐3'
5'‐rArCrCrArCrArArArCrArCrCrArArArUrGrGrUrCrArArCrUrUrGrA‐3'
NC1 Negative Control (Commercial IDT Control):
5'‐rCrGrUrUrArArUrCrGrCrGrUrArUrArArUrArCrGrCrGrUdAdT‐3'
5'‐rArUrArCrGrCrGrUrArUrUrArUrArCrGrCrGrArUrUrArArCrGrArC‐3'
For analysis of Golgi morphology, 10 individual fields positive for EGFP expression were collected from 3 independent
experiments (30 images in total for each condition). All cells in the field were visually scored for Golgi morphology and p125A
expression. Data was analyzed using Windows Excel 2010 (Microsoft Corporation). Homoscedastic two‐tailed Students t‐
tests on percentage of intact Golgi were performed using Windows Excel 2010 (Microsoft Corporation).
Temperature‐block analysis.
HeLa cells were transfected as described above and allowed to express FP‐proteins for 14‐16 hr. Subsequently, cell media
was supplemented with 20 mM HEPES (pH=7.4) (Fisher Scientific) and the cells were incubated at 15°C or 10°C for 4 hr.
Samples were fixed and processed for analysis.
Lipid blot‐overlay
PIP Strips (Echelon) were blocked for 1 hr. in TBS‐Tween buffer (Tris Buffered Saline (TBS), pH=8.0 1% Tween‐20)
supplemented with 3% Bovine Serum Albumin (BSA, Fraction V, EMD) and incubated for 2 hr. in the same buffer
supplemented with 1 μg/mL of GST or His6 tagged proteins. Strips were washed (6 X 5 min. incubations) in TBS‐Tween and
incubated for 1 hr. in TBS‐Tween supplemented with 3 % BSA and HRP‐conjugated murine anti His6 or GST antibodies.
Subsequently, strips were washed (6 X 5 min.) in TBS‐Tween and visualized using SuperSignal West Dura Extended Duration
Substrate (Thermo Scientific) and HyBlot CL X‐Ray film (Denville Scientific Inc.) according to provided protocol.

Immunofluorescence
107
Indirect immunofluorescence was carried out as previously described (Aridor et al., 1995). Images were acquired on an
Olympus Fluoview 1000 confocal system using an inverted microsope (IX‐81 Olympus) and 60X NA 1.42 PLAPON objective.
Images were processed using FV10‐ASW V. 02.00.03.10 (Olympus Corporation) and Adobe Photoshop CS3 (Adobe Photoshop
Version: 10.0.1 (Adobe).
Super‐resolution Structured illumination microscopy was performed on a N‐SIM system (Nikon Instruments, Inc, Melville
NY.) coupled to an inverted microscope (Ti‐E). Z series (0.125 um z steps) were collected with a 100X NA 1.49 Apochromat
total internal reflection oil immersion objective lens. The images were processed with NIS‐Elements software (Nikon
Instruments, Inc, Melville NY) using the following reconstruction parameters: Structured illumination contrast = 2.5,
apodization filter = 0.15, width of 3D‐SIM filter = 0.20 or 0.25. Following reconstruction the data sets were ported to Imaris
(Bitplane) for subsequent processing. Surface rendered volumes were generated using a seed size of 0.1um with a surface
resolution of 0.05 microns. Movies were generated within Imaris, and individual representative frames selected to make
multipanel montages.
Fluorescence Recovery After Photobleaching (FRAP)
HeLa cells were plated at a density of 2 x 10 5 on microwell dishes (35mm) with a 14mm Coverglass bottom No. 1.5 0.16‐0.19
mm (MatTek Corporation) and incubated for 24 hr. Cells were subsequently transfected as described above and incubated
for additional 14‐18 hr. For analysis, cells were washed with PBS and incubated in Phenol‐Red free DMEM (HyClone Fisher‐
Scientific) supplemented with 10 % FBS, 2 mM L‐Glutamine (Gibco‐Invitrogen), 1 mM Na‐Pyruvate (Hyclone, Fisher Scientific),
20 mM HEPES (pH=7.4)(Calbiochem) and 2 % Oxy‐Fluor (Oxyrase). Cells were imaged at 37°C with a PLAPON 60 x objective,
NA = 1.42 at Sampling speed of 10 μs/pixel using an Olympus Fluoview 1000 confocal system. Imaged objects were adjusted
for minimal pixel saturation prior to recording. 5 pre‐bleach reference images were collected prior to photobleaching, which
was achieved by illumination using both 405 and 465 nm lasers for 900ms (100% power). Recovery was recorded in
subsequent 75 to 100 images at 980 ms intervals. Each collected (bleached) region of interest (ROI) (reported as pixel
intensity average) was divided by an adjacent ROI in the same image field (from a non‐bleached spot) to adjust for
background fluctuations and normalized to pre bleached intensity using the average intensity measured in the 5 pre‐
bleached images. Images and ROI's were processed using FV10‐ASW V. 02.00.03.10 software package (Olympus Corporation)
and Windows Excel 2007 (Microsoft Corporation). Data sets from individual experiments were averaged and the values of
the initial 20‐25s time points post‐bleaching were verified for normal distribution using Shapiro‐Wilk (SW) test (p > 0.1) in
Wolfram Mathematica 8 (Wolfram Research). Collected sets of recordings were fitted using Wolfram Mathematica 8
(Wolfram Reasearch) with a reaction‐limited recovery function after adjusting t=0 to the first recorded image post‐bleaching,

108
as described in Forster et al (2006). The presented T1/2 (ln2/k) is an average of T1/2 from fittings of three different sets of
recordings.
Zn 2+ based polymerization assay
GST‐SAM or GST‐SAM L690E were diluted to a final concentration of 10 μM in 50 mM Tris‐HCl (pH=7.5), 100 mM NaCl. An equal
volume of 20 μM Zn(OAc)2 in the same buffer was added and polymerization was estimated by centrifugation at 15000 rpm
using a cooled microfuge (Sorvall). Supernatant and pellet fractions were separated on SDS‐PAGE gels and proteins were
visualized using coomassie blue staining. For thrombin cleaved proteins SAM and SAM L690E and Zn(OAc)2 (both at 0.455 mM)
were incubated in buffer containing 50mM Tris‐HCl (pH=7.5) 100 mM NaCl and polymerization was determined as above.
Proteins were visualized using SilverQuest Staining Kit (Invitrogen).
Coat recruitment to LUVs or ER microsomes
Floatation assays using cytosol, Sar1 proteins and large unilamellar vesicles (LUVs) were performed as previously described
(Bielli et al., 2005). LUVs composition was (Mol / %) 35 % L‐α‐phosphatidylcholine (PC Chicken Egg, Avanti, Polar Lipids, Inc.),
35 % PE (Avanti, Polar Lipids, Inc.), 10 % 1,2‐Dioleoyl‐sn‐glycero‐3‐phospho‐L‐serine sodium salt (PS, Sigma‐Aldrich), 10%
cholesterol (Avanti, Polar Lipids, Inc.), and 10 % L‐α‐phosphatidylinositol‐4‐phosphate (PI4P, Brain, Porcine‐Di ammonium
Salt, Avanti, Polar Lipids, Inc.) or control LUVs containing 45 % PC, 35 % PE, 10% PS, 10 % Cholesterol. For immunodepletion,
anti p125A or control antibodies (anti sorting nexin 9 antibodies kindly provided by Dr. Linton Traub, University of Pittsburgh,
Pittsburgh PA) were used to immunoprecipitate p125A from rat liver cytosol. The resulting supernatants were adjusted for
protein concentration, verified for similar Sec23 and HSP70 levels using western blots and used in microsome binding assays.
Sar1 induced recruitment of Sec23 to ER microsomes was performed as previously described (Pathre et al., 2003).
Endoglycosidase H digestion and analysis of VSV‐G glycosylation
HeLa cells were seeded at 5x10 5 cells/ml on the day before transfection with ts045‐VSV‐G‐Venus and pmRFP‐C1H, pmRFP‐
p125, pmRFP‐p125 PIX/L690E , or pmRFP‐p125 ∆DDHD/L690E using Lipofectamine 2000 (Invitrogen Life Technologies) following
standard protocol. At 6 hours post transfection, cells were shifted to the restrictive temperature of 42°C overnight to block
VSVG in the ER. At 24 hours post‐transfection, cells were treated with fresh media containing 100 μg/mL cycloheximide and
shifted to the permissive temperature of 32°C. At specified time points, cells were collected in PBS containing 1mM DTT and
1mM PMSF by centrifugation at 4°C, resuspended in 100 μL of Endo‐H Buffer (50 mM sodium citrate (pH 5.5), 0.5% SDS, 40
mM DTT) and lysed at 4°C for 30 minutes. Lysates were boiled for 10 minutes, pelleted at 15000 rpm for 15 minutes at 4°C,
and the supernatant transferred to fresh tubes. 25 μL of lysate was diluted 1:1 with Endo‐H Buffer and digested with 2 μL of
Endo H (500 U/ μL, New England BioLabs) at 37°C overnight. VSV‐G was resolved by 8 % SDS‐PAGE and Endo H resistant and

109
sensitive forms of the protein were detected by Western blot analysis utilizing a monoclonal antibody to GFP (Invitrogen
Clone C163).
Acknowledgement
We thank Drs K. Tani, M. Tagaya (Tokyo University, Japan), Dr. W. E. Balch (TSRI, CA), Dr. V Malhotra (GRC, Spain) and Dr. A.
Linstedt (CMU, PA) for valuable reagents. The study was supported by NIH grants 2R56DK0623181 and R01DK092807 (MA).
Abbreviations
Knockdown (KD), Phosphatidylcholine (PC), Phosphatidylinosotol 4‐phosphate (PI4P), Phosphatidic acid (PA),
Phosphatidylethanolamine (PE), phospholipase A1 (PLA1), Endoplasmic reticulum exit sites (ERES), coat protein complex II
(COPII), Vesicular Stomatitis Virus Glycoprotein (VSV‐G).

References
110
Antonny, B., D. Madden, S. Hamamoto, L. Orci, and R. Schekman. 2001. Dynamics of the COPII coat with GTP and stable
analogues. Nat Cell Biol. 3:531‐7.
Antonny, B., and R. Schekman. 2001. ER export: public transportation by the COPII coach. Curr Opin Cell Biol. 13:438‐43.
Aridor, M., K.N. Fish, S. Bannykh, J. Weissman, T.H. Roberts, J. Lippincott‐Schwartz, and W.E. Balch. 2001. The Sar1 GTPase
coordinates biosynthetic cargo selection with endoplasmic reticulum export site assembly. J Cell Biol. 152:213‐29.
Aridor, M., B. S.I., T. Rowe, and W.E. Balch. 1995. Sequential Coupling Between COPII and COPI Vesicle Coats in Endoplasmic
Reticulum to Golgi Transport. J. Cell Biol. 131:875‐893.
Aridor, M., J. Weissman, S. Bannykh, C. Nuoffer, and W.E. Balch. 1998. Cargo Selection by the COPII Budding Machinery
during Export from the ER. J. Cell Biol. 141:61‐70.
Arimitsu, N., T. Kogure, T. Baba, K. Nakao, H. Hamamoto, K. Sekimizu, A. Yamamoto, H. Nakanishi, R. Taguchi, M. Tagaya, and
K. Tani. 2011. p125/Sec23‐interacting protein (Sec23ip) is required for spermiogenesis. FEBS Lett. 585:2171‐6.
Audhya, A., M. Foti, and S.D. Emr. 2000. Distinct roles for the yeast phosphatidylinositol 4‐kinases, Stt4p and Pik1p, in
secretion, cell growth, and organelle membrane dynamics. Mol Biol Cell. 11:2673‐89.
Bielli, A., C.J. Haney, G. Gabreski, S.C. Watkins, S.I. Bannykh, and M. Aridor. 2005. Regulation of Sar1 NH2 terminus by GTP
binding and hydrolysis promotes membrane deformation to control COPII vesicle fission. J Cell Biol. 171:919‐24.
Blumental‐Perry, A., C.J. Haney, K.M. Weixel, S.C. Watkins, O.A. Weisz, and M. Aridor. 2006. Phosphatidylinositol 4‐
phosphate formation at ER exit sites regulates ER export. Dev Cell. 11:671‐82.
Farhan, H., M. Weiss, K. Tani, R.J. Kaufman, and H.P. Hauri. 2008. Adaptation of endoplasmic reticulum exit sites to acute
and chronic increases in cargo load. Embo J. 27:2043‐54.
Fath, S., J.D. Mancias, X. Bi, and J. Goldberg. 2007. Structure and organization of coat proteins in the COPII cage. Cell.
129:1325‐36.
Forster, R., M. Weiss, T. Zimmermann, E.G. Reynaud, F. Verissimo, D.J. Stephens, and R. Pepperkok. 2006. Secretory cargo
regulates the turnover of COPII subunits at single ER exit sites. Curr Biol. 16:173‐9.
Frangioni, J.V., and B.G. Neel. 1993. Solubilization and purification of enzymatically active glutathione S‐transferase (pGEX)
fusion proteins. Anal Biochem. 210:179‐87.
Fromme, J.C., M. Ravazzola, S. Hamamoto, M. Al‐Balwi, W. Eyaid, S.A. Boyadjiev, P. Cosson, R. Schekman, and L. Orci. 2007.
The genetic basis of a craniofacial disease provides insight into COPII coat assembly. Dev Cell. 13:623‐34.

111
Hughes, H., A. Budnik, K. Schmidt, K.J. Palmer, J. Mantell, C. Noakes, A. Johnson, D.A. Carter, P. Verkade, P. Watson, and D.J.
Stephens. 2009. Organisation of human ER‐exit sites: requirements for the localisation of Sec16 to transitional ER. J Cell Sci.
122:2924‐34.
Iinuma, T., A. Shiga, K. Nakamoto, B. O'Brien M, M. Aridor, N. Arimitsu, M. Tagaya, and K. Tani. 2007. Mammalian Sec16/p250
plays a role in membrane traffic from the endoplasmic reticulum. J Biol Chem.
Inoue, H., T. Baba, S. Sato, R. Ohtsuki, A. Takemori, T. Watanabe, M. Tagaya, and K. Tani. 2012. Roles of SAM and DDHD
domains in mammalian intracellular phospholipase A1 KIAA0725p. Biochim Biophys Acta. 1823:930‐9.
Knight, M.J., M.K. Joubert, M.L. Plotkowski, J. Kropat, M. Gingery, F. Sakane, S.S. Merchant, and J.U. Bowie. 2010. Zinc binding
drives sheet formation by the SAM domain of diacylglycerol kinase delta. Biochemistry. 49:9667‐76.
Kuehn, M.J., J.M. Herrmann, and R. Schekman. 1998. COPII‐cargo interactions direct protein sorting into ER‐derived transport
vesicles. Nature. 391:187‐90.
Kung, L.F., S. Pagant, E. Futai, J.G. D'Arcangelo, R. Buchanan, J.C. Dittmar, R.J. Reid, R. Rothstein, S. Hamamoto, E.L. Snapp,
R. Schekman, and E.A. Miller. 2011. Sec24p and Sec16p cooperate to regulate the GTP cycle of the COPII coat. EMBO J.
Lee, M.C., L. Orci, S. Hamamoto, E. Futai, M. Ravazzola, and R. Schekman. 2005. Sar1p N‐terminal helix initiates membrane
curvature and completes the fission of a COPII vesicle. Cell. 122:605‐17.
Litvak, V., N. Dahan, S. Ramachandran, H. Sabanay, and S. Lev. 2005. Maintenance of the diacylglycerol level in the Golgi
apparatus by the Nir2 protein is critical for Golgi secretory function. Nat Cell Biol. 7:225‐34.
Long, K.R., Y. Yamamoto, A.L. Baker, S.C. Watkins, C.B. Coyne, J.F. Conway, and M. Aridor. 2010. Sar1 assembly regulates
membrane constriction and ER export. J Cell Biol. 190:115‐28.
Lorente‐Rodriguez, A., and C. Barlowe. 2011. Requirement for Golgi‐localized PI(4)P in fusion of COPII vesicles with Golgi
compartments. Mol Biol Cell. 22:216‐29.
Matsuoka, K., L. Orci, M. Amherdt, S.Y. Bednarek, S. Hamamoto, R. Schekman, and T. Yeung. 1998. COPII‐coated vesicle
formation reconstituted with purified coat proteins and chemically defined liposomes. Cell. 93:263‐75.
McGary, K.L., T.J. Park, J.O. Woods, H.J. Cha, J.B. Wallingford, and E.M. Marcotte. 2010. Systematic discovery of nonobvious
human disease models through orthologous phenotypes. Proc Natl Acad Sci U S A. 107:6544‐9.
Mezzacasa, A., and A. Helenius. 2002. The transitional ER defines a boundary for quality control in the secretion of tsO45
VSV glycoprotein. Traffic. 3:833‐49.

112
Miller, E.A., T.H. Beilharz, P.N. Malkus, M.C. Lee, S. Hamamoto, L. Orci, and R. Schekman. 2003. Multiple cargo binding sites
on the COPII subunit Sec24p ensure capture of diverse membrane proteins into transport vesicles. Cell. 114:497‐509.
Mizoguchi, T., K. Nakajima, K. Hatsuzawa, M. Nagahama, H.P. Hauri, M. Tagaya, and K. Tani. 2000. Determination of
functional regions of p125, a novel mammalian Sec23p‐ interacting protein. Biochem Biophys Res Commun. 279:144‐9.
Nagaya, H., I. Wada, Y.J. Jia, and H. Kanoh. 2002a. Diacylglycerol kinase delta suppresses ER‐to‐Golgi traffic via its SAM and
PH domains. Mol Biol Cell. 13:302‐16.
Nagaya, H., I. Wada, Y.J. Jia, and H. Kanoh. 2002b. Diacylglycerol Kinase delta Suppresses ER‐to‐Golgi Traffic via Its SAM and
PH Domains. Mol Biol Cell. 13:302‐16.
Nakajima, K., H. Sonoda, T. Mizoguchi, J. Aoki, H. Arai, M. Nagahama, M. Tagaya, and K. Tani. 2002. A novel phospholipase
A1 with sequence homology to a mammalian Sec23p‐interacting protein, p125. J Biol Chem. 277:11329‐35.
Nakatsu, F., R.M. Perera, L. Lucast, R. Zoncu, J. Domin, F.B. Gertler, D. Toomre, and P. De Camilli. 2010. The inositol 5‐
phosphatase SHIP2 regulates endocytic clathrin‐coated pit dynamics. J Cell Biol. 190:307‐15.
Ong, Y.S., B.L. Tang, L.S. Loo, and W. Hong. 2010. p125A exists as part of the mammalian Sec13/Sec31 COPII subcomplex to
facilitate ER‐Golgi transport. J Cell Biol. 190:331‐45.
Pathre, P., K. Shome, A. Blumental‐Perry, A. Bielli, J.H. Haney, S. Alber, S.C. Watkins, G. Romero, and M. Aridor. 2003.
Activation of Phospholipase D by the Small GTPase Sar1 is Required to support COPII Assembly and ER Export. EMBO J.
22:4059‐4068.
Qiao, F., and J.U. Bowie. 2005. The many faces of SAM. Sci STKE. 2005:re7.
Raaijmakers, J.H., L. Deneubourg, H. Rehmann, J. de Koning, Z. Zhang, S. Krugmann, C. Erneux, and J.L. Bos. 2007. The PI3K
effector Arap3 interacts with the PI(3,4,5)P3 phosphatase SHIP2 in a SAM domain‐dependent manner. Cell Signal. 19:1249‐
57.
Rowe, T., M. Aridor, J.M. McCaffery, H. Plutner, C. Nuoffer, and W.E. Balch. 1996. COPII vesicles derived from mammalian
endoplasmic reticulum microsomes recruit COPI. J Cell Biol. 135:895‐911.
Rowe, T., and W.E. Balch. 1995. Expression and purification of mammalian Sarl. Methods Enzymol. 257:49‐53.
Saraste, J., and K. Svensson. 1991. Distribution of the intermediate elements operating in ER to Golgi transport. J Cell Sci:415‐
30.
Sato, S., H. Inoue, T. Kogure, M. Tagaya, and K. Tani. 2010. Golgi‐localized KIAA0725p regulates membrane trafficking from
the Golgi apparatus to the plasma membrane in mammalian cells. FEBS Lett. 584:4389‐95.

113
Shaywitz, D.A., P.J. Espenshade, R.E. Gimeno, and C.A. Kaiser. 1997. COPII subunit interactions in the assembly of the vesicle
coat. J Biol Chem. 272:25413‐6.
Shimoi, W., I. Ezawa, K. Nakamoto, S. Uesaki, G. Gabreski, M. Aridor, A. Yamamoto, M. Nagahama, M. Tagaya, and K. Tani.
2005. p125 is localized in endoplasmic reticulum exit sites and involved in their organization. J Biol Chem. 280:10141‐8.
Stagg, S.M., P. LaPointe, A. Razvi, C. Gurkan, C.S. Potter, B. Carragher, and W.E. Balch. 2008. Structural basis for cargo
regulation of COPII coat assembly. Cell. 134:474‐84.
Supek, F., D.T. Madden, S. Hamamoto, L. Orci, and R. Schekman. 2002. Sec16p potentiates the action of COPII proteins to
bud transport vesicles. J Cell Biol. 158:1029‐38.
Tani, K., T. Mizoguchi, A. Iwamatsu, K. Hatsuzawa, and M. Tagaya. 1999. p125 is a novel mammalian Sec23p‐interacting
protein with structural similarity to phospholipid‐modifying proteins. J Biol Chem. 274:20505‐12.
Weixel, K.M., A. Blumental‐Perry, S.C. Watkins, M. Aridor, and O.A. Weisz. 2005. Distinct Golgi populations of
phosphatidylinositol 4‐phosphate regulated by phosphatidylinositol 4‐kinases. J Biol Chem. 280:10501‐8.
Whittle, J.R., and T.U. Schwartz. 2010. Structure of the Sec13‐Sec16 edge element, a template for assembly of the COPII
vesicle coat. J Cell Biol. 190:347‐61.
Yamashita, A., T. Kumazawa, H. Koga, N. Suzuki, S. Oka, and T. Sugiura. 2010. Generation of lysophosphatidylinositol by DDHD
domain containing 1 (DDHD1): Possible involvement of phospholipase D/phosphatidic acid in the activation of DDHD1.
Biochim Biophys Acta. 1801:711‐20.
Yorimitsu, T., and K. Sato. 2012. Insights into structural and regulatory roles of Sec16 in COPII vesicle formation at ER exit
sites. Mol Biol Cell. 23:2930‐42.
Zanetti, G., K.B. Pahuja, S. Studer, S. Shim, and R. Schekman. 2011. COPII and the regulation of protein sorting in mammals.
Nat Cell Biol. 14:20‐8.

Figure legends:
114
Fig. 1. Sar1 dependent COPII and p125A recruitment requires PI4P.
A. Sar1 dependent Sec23 recruitment to floated liposomes is dependent on PI4P. Active
(Sar1A‐GTP) or inactive (Sar1A‐GDP, both tested at 1 g, 50 L final volume) were incubated
with rat liver cytosol and synthetic Large Unilammear Vesicles (LUV, 400 M) composed of 45
% PC, 35 % PE, 10% PS, and 10 % Cholesterol or 35 % PC, 35 % PE, 10% PS, 10 % Cholesterol,
10 % PI4P at 26°C for 1 hr. and floated onto a sucrose gradient. Fractions were analyzed by
western blot with antibodies against Sec23. (Floated fractions as labeled). B. Sec23
recruitment to ER membrane is not affected by depletion of p125A. p125A or SNX9 (control)
were depleted from rat liver cytosol by immunoprecipitation and p125A depletion was
verified using western blot as indicated. Sar1A‐GTP dependent Sec23 recruitment from
control or p125A depleted cytosol to ER microsomes was monitored. Sec23 was recruited by
Sar1A‐GTP (50 ng, 500 ng and 1 g, final volume 60 L) in a dose dependent manner in the
absence (lanes 1‐4) or presence (lanes 5‐8) of p125A. C. Co‐recruitment of Sec23 and p125A
to synthetic PI4P containing LUVs. Synthetic LUV's composed by 35 % PC, 35 % PE, 10% PS, 10
% Cholesterol, and 10 % PI4P were incubated and fractionated as in A. in the presence of Sar1‐
GTP or Sar1‐GDP as indicated. Sec23, Sec31a and p125A recruitment (as indicated) to floated
liposomes was monitored by western blot.
Fig. 2. p125A is targeted to ER exit sites whereas isolated DDHD domain is targeted to Golgi
membranes.
A. Transiently expressed EGFP tagged p125A co‐localizes predominantly with Sec31 (as
observed with the endogenous p125A protein) at ERES and was juxtaposed to ERGIC

115
(ERGIC53) or cis‐Golgi compartments (GPP130). B. Transiently expressed EGFP tagged DDHD
domain dissociated from Sec31 containing ERES (left image arrowhead shows lack of co‐
localization between Sec31 stained ERES and localized EGFP‐DDHD). The DDHD domain
targets to the periphery of PI4P enriched membranes and can be seen coating Golgi (right
image, TGN46) and ERGIC (ERGIC 53, middle image). Bars in all figures are 10m.
Fig. 3. Cooperative lipid recognition by the SAM‐DDHD module of p125A.
A. Structural Model of p125A‐Sec23 interactions. p125A consists of a proline‐glutamine(P‐Q)
rich N‐terminus that contains a WWE domain (thin lined boxes). The C‐terminus of p125A
consists of DDHD domain and a SAM domain (marked in bold box). Arrowhead indicates the
position of leu 690 in a structured model of p125A’s SAM domain (see also panel G). The
structure of the Sar1 (green)‐Sec23 (blue) complex assembled with the Sec31A active peptide
(pink) is shown (PDB 2QTV) where p125A is predicted to interact with Sec23 using the N‐
terminus P‐Q domain. B. The DDHD domain targets PI4P rich Golgi membrane in isolation.
Transiently expressed GFP tagged DDHD domain (779‐989) is targeted to PI4P enriched Golgi
membranes (left image). Replacing a basic stretch of residues in the DDHD domain with
glutamic acid residues (850‐KGRKR‐854EGEEE, termed PI‐X) abolished Golgi targeting (right
image) C‐E. Selective lipid recognition is dependent on a module consisting of the DDHD and
SAM domains. C. 1 μg/mL of purified His tagged p125A fragment (701‐989) containing the
DDHD domain but not the SAM domain was probed on lipid blot overlay using HRP conjugated
antibody against His6. The 701‐989 fragment bound weakly to phosphorylated
phosphoinositides PA and PS. D. As in C. Extending the fragment to contain the upstream SAM
domain (643‐989) conferred lipid selectivity to monophosphorylated PIs, PA, PS and PI(3,4)P2.
E. As in C. PI‐X mutations in the DDHD domain of the SAM‐DDHD module abolished lipid

116
recognition F. The EGFP‐SAM domain transiently expressed in HeLa cells shows diffused
cytosolic distribution. G. The predicted structure of p125A’s SAM domain (generated in
Phyre). The red arrowhead indicates the position of a conserved leucine (690) within a
predicted hydrophobic dimer interface. H. GST‐SAM oligomerization is promoted by Zinc
addition and is sensitive to the L690E mutation. 20 μM Zn(AOc)2, or buffer as control was
added to GST‐SAM or GST‐SAM (L690E, 10 μM of each) as indicated. Oligomerization was
followed by the precipitation of the proteins from supernatant (S) to pellet (P) fractions (as
indicated) using centrifugation and analysis on coomassie stained gels. I. SAM and SAM L690E
domains (cleaved of GST, 0.455mM each) were incubated with 0.455mM of Zn(AOc)2 and
analyzed as in H.
Fig. 4. ERES coated p125A segregates from ERGIC and Golgi compartments at low
temperatures.
HeLa cells transiently expressing mRFP‐p125A were maintained at 37°C, or incubated at 15°C
or 10°C as indicated for 4 hr., fixed and analyzed for localization with ERES (hSec31A, A).
Bottom panel are enlarged images of boxed areas, arrowheads highlight the extensive co‐
localization of p125A to ERES. Similar analysis shows lack of co‐localization with ERGIC
(ERGIC53, B) or Golgi (gp73, C) as indicated. Note the segregation of ERGIC (B) or Golgi (C)
from p125A coated ERES under these conditions as highlighted by arrowheads in the merged
images.
Fig. 5. ERES coated p125A segregates from ERGIC and Golgi compartments at low
temperatures.

117
A. HeLa cells transiently expressing EGFP‐mSec16A were maintained at 37°C, or incubated at
15°C or 10°C as in Fig. 4 and the localization of ERES (marked by hSec31a) and EGFP‐mSec16A
was determined. B. HeLa cells transiently expressing mRFP‐p125A were incubated at 10°C and
the localization of mRFP‐p125A and endogenous mSec16A was determined.
Fig. 6. PI4P binding by the SAM‐DDHD module of p125A controls ERES assembly.
A. The localization of transiently expressed full‐length EGFP‐p125A wt, p125A PI‐X or L690E
and the double mutant (p125A L690E,PI‐X ) and ERES (hSec31a) was analyzed in HeLa cells as
indicated. The bottom panel shows the expression of a chimera where the DDHD domain has
been substituted with the PH domain from Fapp1 in the backbone of an L690E mutant. EGFP‐
p125API‐X or L690E PI‐X, L690E
mutants become partly cytosolic whereas the double mutant p125A
lost membrane localization and completely disrupted ERES assembly. Membrane targeting
and ERES assembly was partially restored in the Fapp1‐PH containing p125A chimera
expressing cells.
Fig. 7. Lipid recognition by p125A regulates mSec16A displacement from ERES. HeLa cells
transiently expressing mRFP‐p125A (A‐B) or mRFP‐p125A L690E, DDHD (C‐D), GFP‐Sec16A (A) or
YFP‐Sec23A (B) for 24 hr, were fixed and analyzed for the localization of transfected proteins
or hSec31a (C). Note the segregation of GFP‐mSec16A from mRFP‐p125A as opposed to the
assembly of mRFP‐p125A with YFP‐Sec23a (A and B). Note the disassembly of ERES (hSec31a)
in cells expressing mRFP p125A L690E, DDHD ( in C), and the collection of GFP‐mSec16A in these
sites (D). Bar is 10 m.

118
Fig. 8. High‐resolution analysis of p125A induced enlarged ERES. A. Confocal images of cells
expressing high levels of wt, PI‐X‐L690E, or L690E, DDHD, mRFP‐p125A proteins as indicated.
B. SIM reconstruction of cells expressing GFP‐mSec16A with mRFP‐p125A or mRFP‐p125A
L690E, DDHD as indicated, fixed and processed using Imaris. Note the engulfment of ERES by
Sec16 with p125A L690E, DDHD . C. SIM reconstruction of cells expressing Venus‐VSV‐G ts (green)
and mRFP‐p125A (red) at time points following a shift of expressing cells to the permissive
temperature as indicated. Note budding of uncoated structures emerging from sites heavily
coated with p125A. Images were processed using NIS elements. Insert shows western blot of
undigested or Endo‐H digested VSV‐G ts at 0 and 120 minutes in cells expressing Sar1 H79G
(negative control), mRFP (positive control) or mRFP‐p125A as indicated.
Fig. 9. Functional SAM‐DDHD module is required for steady state ER to Golgi traffic.
A. Typical observed Golgi morphologies are shown and color‐coded including Intact (blue),
Dispersed (pink) and Shattered (vesiculated, green) as indicated. In some cells Golgi was not
recognizable (labeled as missing, purple) B. Analysis of p125A knockdown efficiency by
western blots. Endogenous p125A expression of p125A (middle panel) and actin (lower panel)
are shown. The upper panel shows the comparable expression of EGFP‐p125A resistant clones
in control and KD cells. EGFP ran below the analyzed area and is not shown. The expression
of EGFP‐p125A resistant WT and L690E, PI‐X clones was also detected by the p125A specific
antibody but required prolonged exposures due to partial transfection efficiency of KD cells.
C. The fractional distribution of Golgi morphologies in control, p125A depleted and rescued
cell populations with EGFP‐p125A and EGFP‐p125A L690E, PI‐X as indicated. The bar diagram
shows the percentage of cells with either intact Golgi (blue), dispersed Golgi (red), shattered
Golgi (green) or missing Golgi (purple) under each treatment condition. D. Statistical analysis

119
of intact Golgi morphology in control and knockdown cells (means and SD). Three individual
KD experiments were performed for each condition. 10 images were collected from each
experiment and Golgi phenotypes were determined for all cells expressing EGFP tagged
proteins. For controls, cells expressing EGFP (n = 220), EGFP‐p125A (n = 103), EGFP‐p125A
L690E,PI‐X (n = 133) were counted. For RNAi treated cells, EGFP (n = 150), EGFP‐p125A (n = 169),
EGFP‐p125A L690E, PI‐X (n = 129) were counted. Unpaired student t‐test between groups is
shown as indicated (**)
Fig. 10. The COPII budding cascade at ERES. Following initial association of COPII layers with
Sec16 on ER membranes (A), the binding of PI4P by p125A promotes the displacement of
Sec16 from COPII inner and outer layers to allow for effective linking between coat layers,
driving coat retention (B) while enhancing GTP hydrolysis to support vesicle fission (C).

Supplement.
120
Fig. S1. The SAM‐DDHD module controls p125A dynamics at ERES.
A. Individual images of cells expressing YFP‐Sec23 and EGFPp125A taken from a typical FRAP
analysis at time intervals as indicated. Arrowheads point to bleached ERES (sites enlarged in
boxed areas). B. Transiently expressed YFP‐Sec23 or EGFP‐p125A were bleached and the rate
of fluorescence recovery was recorded. Average from 35 individual measurements of
EGFPp125A (C), EYFP‐Sec23 (35 measurements, B), EGFPp125A PI‐X (33 measurements, E) or
EGFPp125A L690E (27 measurements, D) collected in three independent experiments are shown
with standard deviation. Note the slower rate of EGFPp125A when compared to YFP‐Sec23.
Note the faster recovery of EGFPp125A PI‐X or EGFPp125A L690E compared to wt, suggesting that
the SAM‐DDHD module controls the dynamics of p125A at ERES.
Fig. S2. p125A depletion induces massive vesiculation of the Golgi complex.
Hela cells stably expressing GFP tagged N‐acetylgalactosaminyltransferase‐2 (GalNAcT2‐GFP,
kindly provided by Dr. B. Storrie, University of Arkansas) were treated with control or p125A
directed DsRNAi (materials and methods) as indicated and visualized for GFP. Note the
dramatic change in Golgi morphology with loss of intact Golgi populations and the
concomitant increase in shattered Golgi morphology. Bar is 10m.

121
Movie S1. Over expressed mRFP‐p125A induces the formation of enlarged structures with
adjacent mSec16A.
HeLa cells over‐expressing mRFP‐p125A and EGFP‐mSec16A (as in Fig. 8B) were fixed and
visualized using SIM microscopy. Projection images were prepared using Imaris software as
described in materials and methods.
Movie S2. Over expressed mRFP‐p125A L690E, DDHD induces the formation of enlarged
structures engulfed by GFP‐mSec16A.
HeLa cells over‐expressing mRFP‐p125A L690E, DDHD and EGFP‐mSec16A (as in Fig. 8B) were
fixed and visualized using SIM microscopy. Projection images were prepared using Imaris
software as described in materials and methods.
Movie S3‐5 Bud structures containing VSV‐G ts ‐Venus emanating from mRFP coated large
structures at 0’ (Movie 3), 60’ (Movie 4) and 90’ (movie 5). Hela cells over‐expressing mRFP‐
p125A and VSV‐G ts ‐Venus were shifted from a non‐permissive to a permissive temperature
(as described in materials and methods), fixed at the indicated time points following
temperature shift and visualized using SIM microscopy (as in Fig. 8C). Projection images were
prepared using NIS‐elements software as described in materials and methods.

74.1
114.3
114.3
74.1
114.3
114.3
A.
74.1
74.1
74.1
74.1
C.
Floated
fraction
1 2 3 4 5 6 7 8 9 10 11 12
Floated
fraction
Sec23
1 2 3 4 5 6 7 8 9 10 11 12
Sar1-GDP
Sar1-GTP
Sar1-GDP
Sar1-GTP
Sec23
p125A
Sec31a
Sec23
no
PI4P
Plus
PI4P
B.
121
Sar1-GDP
p125A Sar1-GTP
Sec31a
122
p125A
Non depleted
p125A depletion
SNX9 depletion
(control)
121
96
p125A
Depleted
Control
(SNX9 depleted)
Sar1-GTP - -
Depleted cytosol + + + + - - - -
Control depleted - - - - + + + +
Microsomes + + + + + + + +
Fig. 1
Klinkenberg et al.
p125A
Sec23

A. EGFP-p125A (full length)
Sec31 ERGIC53 GPP130
B. EGFP-DDHD domain
123
Sec31 ERGIC53
TGN46
Fig. 2
Klinkenberg et al.

A.
WWE
P-Q
B.
GFP-DDHD GFP-DDHD-(PI-X)
C.
LPA
LPS
PI
PIP(3)
PIP(4)
PIP(5)
PE
PC
DDHD
++++
DDHD
SIP
PIP 2 (3,4)
PIP 2 (3,5)
PIP 2 (4,5)
PIP 3 (3,4,5)
PA
PS
Sam
DDHD
++++
Blank
D. E.
LPA
LPS
PI
PIP(3)
PIP(4)
PIP(5)
PE
PC
Sam-
DDHD
SIP
PIP 2 (3,4)
PIP2 (3,5)
PIP2 (4,5)
PIP3 (3,4,5)
PA
PS
Blank
LPA
LPS
PI
PIP(3)
PIP(4)
PIP(5)
Sam-
DDHD(PI-X)
PE
PC
P-Q
124
SIP
PIP 2 (3,4)
PIP 2 (3,5)
PIP 2 (4,5)
PIP 3 (3,4,5)
PA
PS
Blank
F.
G.
H.
101.5
87.6
GST-
Sam
GST-
SamL690E
P S P S P S P S
Zn ++ + - + -
I.
201.2
114.3
74.1
Zn ++
52.7
36.7
27.8
18.8
48
34.4
27.2
17.0
6.4
GFP-Sam
Sam
Sam L690E
P S P S
+ +
Fig. 3
Klinkenberg et al.

A. ERES
37°C
15°C
10°C
125
hSec31a mRFP-p125A Merge
37°C 15°C 10°C
Fig. 4
Klinkenberg et al.

B. ERGIC
C. Golgi
37°C
15°C
10°C
15°C
10°C
ERGIC53
gp73
126
mRFP-p125A Merge
mRFP-p125A Merge
Fig. 4B-C
Klinkenberg et al.

37°C
15°C
10°C
10°C
EGFP-mSec16A hSec31a Merge
127
mRFP-p125A mSec16A (KIAA0310) Merge
Fig. 5
Klinkenberg et al.

WT
PI-X
EGFP-p125A hSec31a Merge
L690E
PI-X, L690E
ΔDDDHD, L690E
+Fapp1-PH
128
* * *
Fig. 6
Klinkenberg et al.

A.
B.
C.
D.
EGFP-mSec16A mRFP-p125A
Merge
YFP-Sec23a mRFP-p125A
Merge
hSec31a
EGFP-mSec16A
129
mRFP-p125A L690E,ΔDDHD
Merge
* * *
mRFP-p125A L690E,ΔDDHD
Merge
Fig. 7
Klinkenberg et al.

A.
WT PI-X, L690E ΔDDDHD, L690E
B. p125A
C.
mSec16A
WT ΔDDHD, L690E
p125A
VSV-G
0' 60' 90'
Undigested
R
S
130
WT ΔDDHD, L690E
Sar1 H79G
0' 120'
mRFP mRFPp125A
0' 120' 0' 120'
Fig. 8
Klinkenberg et al.

A.
B.
EGFP
p125A
Actin
Intact Dispersed Shattered
Control RNAi
1 2 3 4 5 6 7 8
EGFP
EGFP-p125A
EGFP-p125AL690E, PI-X
EGFP
EGFP-p125A
EGFP-p125AL690E, PI-X
114.3
114.3
48
C.
EGFP
EGFP-p125A
131
Control RNAi
EGFP-p125AL690E, PI-X
EGFP
EGFP-p125A
EGFP-p125AL690E, PI-X
D.
% cells with intact Golgi
50
40
30
20
10
EGFP
EGFP-p125A
Control RNAi
EGFP-p125AL690E, PI-X
Fig. 9
Klinkenberg et al
Missing
Shattered
Dispersed
Intact
**
P

A. B. C.
NHH 2
p125A
mSec16A
Sec23/Sec24
Sec13/Sec31
Bet3-TRAPPI
Sar1-GTP
Sar1-GDP
PI4P
CCOOHH
NNHH 22
132
CCOOOOH
Fig. 10
Klinkenberg et al.

A.
1.2
1
0.8
0.6
0.4
0.2
YFP-Sec23a
EGFP-p125A
133
B. C. D.
YFP-Sec23a EGFP-p125A
EGFP-p125AL690E EGFP-p125API-X 0
0 50 100 150 0 50 100 150 0 50 100 150 0 50 100 150
(Sec.)
E.
Fig. S1
Klinkenberg et al.

134
Control RNAi
Fig. S2
Klinkenberg et al.

Investigations of p125A‐Sec31A associations
and mammalian Sec16A and B membrane
binding
Additional exploration of p125A
The SAM (L690) or DDHD (PI‐X) mutations abrogate Golgi targeting of p125A (643‐989)
135
We wished to further explore the targeting towards PI(4)P of the SAM and the DDHD
domain in the p125A (643‐989) fragment in in vivo settings. For this purpose mRFP‐tagged
versions of wt and mutant forms of the SAM and the DDHD domain fragments were cloned
(see fig 1). The fragments were transiently transfected into HeLa cells and their localization
were examined by fluorescent confocal microscopy.
Figure 1 – Graphical overview of p125A ‐ The L690E mutation in the SAM domain and the 850
KGRKR/EGEEE 854 (PI‐X) mutation in the DDHD domain are depicted.
Analysis of the cellular localization of the mRFP‐tagged p125A (643‐989) fragment
containing the wt SAM and DDHD domains during low level overexpression showed
targeting to PI(4)P enriched membranes of both the Golgi and ERES (see fig. 2A). During high
level overexpression, the fragment showed a tendency to aggregate in larger structures that
frequently perturbed Sec31A distribution (data not shown). Sec31A also tended to collect in
aggregates next to the p125A (643‐989) during high level over‐expression of the fragment,
and these cells regularly exhibited a dispersed Golgi.
Introducing the DDHD (PI‐X) mutation to the p125A (643‐989) fragment caused a clear loss
of membrane association resulting in a predominant cytosolic distribution of the fragment
(see fig 2B). Introducing both the SAM(L690E) mutation and the (PI‐X) mutation to the
fragment also exhibited cytosolic distribution without specific targeting (see fig. 2D). Sec31A
distribution was generally perturbed in p125A (643‐989)(L690E)(PI‐X) expressing cells.

136
Occasionally during high levels of overexpression of p125A (643‐989)(L690E)(PI‐X), Sec31A
was observed collect in larger aggregate structures that co‐localized with the p125A (643‐
989)(L690E)(PI‐X) fragment.
Figure 1 – p125A (643‐989) wt and mutant fragments are expressed predominantly as cytosolic proteins – All four
variants of mRFPp125A (643‐989) were transiently expressed in HeLa cells, fixed in 3.7 % formaldehyde solution and co‐
stained with ERES specific antibodies raised against Sec31A (green) and cis‐Golgi marker GPP73 (white). A) mRFP‐tagged wt
p125A (643‐989) targeted towards the PI(4)P rich membranes of Golgi and ERES, with an appreciable cytosolic background
stain. B & C) Introduction of either the PI‐X mutation (inhibiting lipid recognition) or the L690E (inhibiting SAM
oligomerization) caused loss of targeting towards Golgi and ERES. D) Expression of the double mutant p125A (643‐
989)(L690E)(PI‐X) resulted in a predominantly cytosolic distribution. Occasionally the p125A (643‐989)(L690E)(PI‐X)
fragment did aggregate into punctate structures that co‐localized with Sec31A. More noticable was the perturbation of the
Sec31A distribution (compare the two cells marked with asterisks). Sec31A would intermittently collect into larger
aggregates that showed some co‐localization with p125A(643‐989)(L690E)(PI‐X) punctae (see box).
Surprisingly, introduction of only the (L690E) mutation caused the fragment to exhibit a
predominantly cytosolic distribution (see fig 2C). Rarely did we observe aggregation and co‐
localization to either ERES or Golgi during high levels of over‐expression (data not shown).
These observations were puzzling, given our observation that expression of the DDHD

137
domain alone showed a high degree of targeting towards PI(4)P‐enriched membranes and in
particular the Golgi.
The reason for the p125A (643‐989)(L690E) loss of targeting was not fully resolved, but
could be explained if the mutation of the SAM domain causes an "inhibitory fold" that
shields the DDHD from binding to lipids. Un‐shielding of the DDHD domain would occur
through the SAM homo‐dimerization, which thereby promotes DDHD targeting to charged
lipids and in particular PI(4)P. Introduction of the L690E mutation inhibts SAM domain
dimerization. Thereby, the mutation maintains the fragment in a DDHD shielded
conformation abbrogating the lipid targeting. Taken together, these result further suggest
that the oligomerization of SAM domains modulates the affinity and avidity of the DDHD
domain towards PI(4)P‐enriched ERES membranes.
EGFPp125A proline‐glutamine (P‐Q) rich region forms large non‐dynamic aggregates
containing Sec31A
We further wished to explore the potential binding of p125A with Sec31A. For these studies
the Proline‐Glutamine (P‐Q) rich N‐terminus of p125A (31‐310) was isolated and cloned into
an EGFP expression vector. Upon transient expression in HeLa cells, EGFP‐tagged p125A (P‐
Q) aggregated into large distinct structures. Staining against Sec31A showed strong
localization to these structures indicating association of Sec31A to the p125A (P‐Q) rich
region (see fig 3A). We were also able detect Sec23 and Sec16 co‐localizing to the p125A (P‐
Q) aggregates (data not shown), which supports that the P‐Q rich region provides binding
sites for Sec23 and Sec31A.
Earlier studies have shown that p125A associates with Sec31A through a region comprising
of the p125A residues 259‐600 [1]. Our results show that p125A region comprising of 31‐
310 associates with Sec31A. Taken together our results imply that p125A binds to Sec31A
through the region of residues 259‐310. Moreover, it is known that the stretch containing
residues 135‐259 of p125A interacts with Sec23 [2]. Both these stretches reside within the
P‐Q rich region of p125A, implying that the P‐Q rich region mediates COPII binding and
linkage.

138
FRAP analysis of the dynamics of EGFPp125A (P‐Q) when collected at the aggregate
structures showed that these structures were rather static with minimal or no exchange of
EGFPp125A (P‐Q) occurring (see fig 3B). This result implies that expression of the p125A (P‐
Q) domain alone sequesters Sec31A and likely also Sec23 into non‐dynamic aggregates.
Identification of a WWE domain in the p125A (P‐Q) region
Figure 3 ‐ p125A (P‐Q) Co‐localization and dynamics ‐ A) EGFPp125A (P‐Q)
was expressed transiently in HeLa cells, fixed and co‐stained with
antibodies raised against Sec31A for ERES localization markers (red). p125A
and Sec31A co‐localize in large aggregate structures in the cells, which can
be seen more clearly in the enlargement window. B) p125A (P‐Q) dynamics
by FRAP. Bleaching of EGFPp125A (P‐Q) shows minimal to no recovery.
Average from 16 recordings with standard deviation.
In order to investigate whether a Sec31 binding module could be within residues 259‐310
we analyzed the p125A N‐terminus with the (P‐Q) rich region for distinct structural features.
The sequence‐based analysis was done using an online Protein Homology/analogy
Recognition Engine V 2.0 (Phyre2 ‐ http://www.sbg.bio.ic.ac.uk/phyre2). The analysis
showed strong structural folding homology in residues 259‐342 (98.5 % confidence) to a
motif regularly associated with E3 ligases and poly‐ADP‐ribose polymerases. This motif is
named WWE after a set of characteristic conserved tryptophan‐tryptophan‐glutamic acid

139
(WWE) residues present (see fig. 4) [3, 4]. A homologous WWE motif has recently also been
identified in p125B [5].
mRFPp125A WWE (259‐342) association with Sec31A and cellular localization
It has previously been shown that p125A maintains association with Sec31A in the cytosol
when not bound to membranes [1]. These observations imply that p125A and Sec31A co‐
exist in a stable complex, and are recruited together to ERES.
Figure 4 ‐ Modeled structure of the
p125A WWE domain ‐ Structure of
WWE domain obtained from structure
alignment of the p125A (P‐Q) fragment
through Phyre2 (Structure ID: D1UJRA).
The WWE domain consists of 6 β‐
strands that form a single twisted anti‐
parallel β‐sheet (yellow) that cups
towards a single 3 turn α‐helix (pink).
Blue regions represent modeled turns
whereas white regions represent
predicted non‐structured residue
stretches.
We hypothesized that p125A might associate with Sec31A by binding to its unstructured
region using the WWE motif. We reasoned that the actual binding of Sec31A to the motif
would ocur within the initial 50 residues of the p125A WWE, and that the Sec31A binding to
p125A WWE would be stabilized by further association with Sec23. This would explain the
Sec31A sequestration observed when expressing the p125A (P‐Q) fragment, as this fragment
encompasses the first 50 residues of the WWE domain as well as the binding site for Sec23.
The identified p125A WWE motif (259‐342) was cloned into a mammalian expression vector
fused to mRFP. The p125A WWE was expressed uniformly throughout transiently
transfected HeLa cells (see fig 5A), with no visible effect on Sec31A‐marked ERES
distribution. At very high expression levels, p125A WWE had a tendency to aggregate and
collect Sec31A in larger distinct punctae (see fig 5B). This observation suggests that p125A

WWE may influence Sec31A in vivo. Co‐localization between p125A (WWE) and Sec31A
could not be observed.
140
We have purified and tested p125A WWE fragment in initial GST pull‐down experiments.
The results thereof need to be further examined and verified.
Figure 5 ‐ mRFPp125A WWE expression and localization ‐ A) mRFPp125A WWE (red) was expressed in HeLa cells, fixed and
co‐stained with an ERES specific antibody raised against Sec31A (green). Normal levels of EGFPp125A WWE expresses
throughout the entire cell with slight reticulation and no apparent influence upon Sec31A expression and ERES distribution.
B) High expression levels of p125A WWE causes both p125A WWE and Sec31A to aggregate into larger puncta and decreases
visible ERES distribution. p125A WWE and Sec31A do not co‐localize strongly.

A study of Sec16A and B membrane binding
Initial aim of the Study
We set out to explore mechanisms that possibly couple the generation of selective lipid
141
signals on ER membranes with the assembly of COPII at ERES. We furthermore also wished
to couple the generation of selective lipid signals with the regulation of ER export. Such
regulation has previously been demonstrated, but the molecular basis remains undefined
(see Introduction). As candidates for mediators of such regulation we focused on Sec16 and
p125A, both being previously shown to directly bind COPII subunits [1, 2, 5‐12].
In this part of the project we hypothesized that Sec16A and Sec16B are accessory proteins
that regulate ERES assembly in response to lipid signaling. We sought to explore this
hypothesis by examining regions of Sec16A and Sec16B that were previously reported to
facilitate ERES targeting, and potentially membrane targeting [13, 14]. This was done by
examining wild type Sec16A and B localization at 37°C, 15°C and 10°C by confocal imaging.
Moreover, Sec16A and Sec16B fragments previously suggested to be sufficient for targeting
to ERES were purified as GST‐tagged fragments. These fragment were examined for Sar1‐
dependent interactions and recruitment to ER microsomes. The same fragments were also
examined for their function in ERES assembly [13, 14].
EGFP‐Sec16A and EGFP‐Sec16B localization
We started out investigating the localization of Sec16 with emphasis on presumed
associations with ERES and markers of the early biosynthetic transport pathway. EGFP‐
tagged Sec16A and Sec16B (kindly provided by Dr. Vivek Malhotra) were transiently
expressed in HeLa cells and analyzed by indirect immunofluorescence. At low expression
levels, EGFP‐Sec16A was distributed throughout the cells in a uniform dispersed pattern
likely covering both ER membranes and the cytosol ‐ the protein often localized at specific
punctae adjacent to and often overlapping with ERES marked by Sec31A (see fig 6A). At
high expression levels, EGFP‐Sec16A maintained the uniform dispersed pattern, whereas the
amount of defined punctae decreased, both for Sec16A and for Sec31A‐marked ERES.
Apparent co‐localization between remaining Sec16A punctae and ERES was still observed
(see fig 6B).

142
In contrast, EGFP‐Sec16B showed a more distinct punctate distribution at low level
expression where several punctae localized and overlapped with Sec31A staining, indicating
association at ERES (see fig 6C). Overexpression resulted in similar uniform dispersed
distribution as observed for Sec16A, and Sec31A also exhibited a more dispersed
distribution (see fig 6D). These results corroborated previous reports of both Sec16A and
Sec16B partially localizing to COPII‐coated ERES [13, 14].
Figure 6 ‐ EGFP‐Sec16A & B expression and cellular localization (green) ‐ EGFP‐Sec16A & EGFP‐Sec16B were
expressed in HeLa cells, fixed and co‐stained with ERES specific antibodies raised against Sec31A (red). A) Low level
EGFP‐Sec16A expression shows dispersed distribution of the protein throughout the cell with defined punctae that are
adjacent and overlap with Sec31A marked ERES. B) High expression of EGFP‐Sec16A shows a similar dispersion with
less apparent Sec16A (green) punctae visible. A similar reduction in Sec31A stained ERES (red) is also observed, while
overlap in localization between Sec16A punctae and Sec31A marked ERES is maintained. C) At low level expression
EGFP‐Sec16B is visible as distinct punctae that in several instances are adjacent and overlap with Sec31A marked ERES.
D) At high level expression, EGFP‐Sec16B becomes more diffuse throughout the cell, and Sec31A becomes noticeably
dispersed when compared to surrounding cells (arrows).

143
EGFP‐Sec16B associates with Sec31A at 37°C, but aggregates into separate structures from
Sec31A, ERGIC53 and Golgi at 15°C and 10°C.
We utilized temperature‐dependent blocking of traffic between ER and the Golgi to examine
the association between Sec16B and ERES. Cells transiently expressing EGFP‐Sec16B were
maintained at 37°C then shifted to lower temperature for 4 h. Here we used either 15°C to
arrest transport at the ERGIC [15], or 10 °C to arrest transport at the ERES (see fig 7A and B)
[16]. At 37°C (see fig 7B), EGFP‐Sec16B expression was either uniform throughout the cell
suggesting a cytosolic distribution, or visible in punctate structures that localized mostly
near Sec31A, and to a lesser extent ERGIC53. EGFP‐Sec16B did not co‐localize with GPP73‐
stained Golgi compartments. Sec31A staining was predominantly dispersed in cells that
overexpressed EGFP‐Sec16B.
At 15°C (see fig 7B), EGFP‐Sec16B organized in defined larger structures that clearly
segregated from Sec31A, ERGIC 53 and GPP73, although the structures clustered in close
vicinity or adjacent to both ERGIC53 and GPP73 containing compartments. This organization
was kept at 10°C (see fig 7A and B). These results imply that Sec16B might be involved in
early stage ERES assembly that can be disconnected from late COPII‐mediated budding. The
results are in agreement with recent observation made by Hughes H. et al [17], showing a
clear spatial separation between Sec16 and Sec31 in C. Elegans.
Figure 7 – A) (Above) EGFP‐Sec16 B expression and localization in HeLa cells at 10°C ‐ EGFP‐Sec16B expressing HeLa cells
incubated at 10°C. Samples were fixed and stained against ERES and cis‐Golgi as described for figure 6A. EGFP‐Sec16B
(green) assembles into larger punctae that separate from Sec31A marked ERES (red) and clusters in the vicinity of GPP73
marked Golgi compartments (white) (arrow). B ) (Next page) EGFP‐Sec16B expression and localization at 37°C and low
temperatures ‐ EGFP‐Sec16B expressing HeLa cells were maintained at 37°C (upper panel), or incubated at 15°C (middle
panel), or 10°C (lower panel). Samples were fixed and stained against ERES with antibodies raised against Sec31A, cis‐Golgi
with antibodies raised against GPP73 and ERGIC with antibodies raised against ERGIC53. At 37°C the EGFP‐Sec16B (green)
appears uniformly distributed throughout the cell with low expression showing puncta that localize adjacent to Sec31A (red).
At 15°C and 10°C the protein clustered in larger puncta, and formed larger structures in vicinity of ERGIC (red) and Golgi
(white) (see magnification window).

144

145
GST‐Sec16B (35‐194) associates with purified NRK microsomes independently of Sar1A
activation and COPII recruitment
For analysis of the molecular basis for Sec16A and B targeting to membranes and in
particular ERES, we were guided by previous findings of Bhattacharyya D. & Glick B.S. [13].
They have defined minimal Sec16A and Sec16B fragments essential and sufficient for ERES
targeting. These studies showed that Sec16B associates with tER sites through an N‐terminal
domain that does not contain the CCD. EGFP‐tagged hybrid fragments connected to the
sequence upstream of the CCD from residue 34 to 234 showed tER localization when
expressed in HeLa cells. Truncation of the N‐terminal 70 residues (and further) caused loss
of tER localization, as did deletion of the 34‐234 segment [13].
Using Sec16B cDNA, a fragment comprising of residues 35‐194 was cloned into pGEX‐4T‐1,
expressed in E. coli BL21 strain, and purified as a GST fusion protein. Our initial aim was to
clone the fragment studied by Bhattacharyya and Glick comprising residues 35‐234 for in
vitro analysis. As a variation at position 195 (arginine to glutamine) was found in our cDNA,
we decided to begin by analyzing a fragment that terminated at this position.
An established COPII recruitment assay was used to examine recruitment of the GST‐Sec16B
(35‐194) to ER membranes. ER microsomes derived from NRK cells were incubated with rat
liver cytosol (RLC) and the active form of Sar1A (H79G). Membranes were collected by
centrifugation and analyzed by Western blotting [18].
GST‐Sec16B (35‐194) bound ER microsomes effectively and independent of Sar1A activation
(see fig 8, lanes 5‐7). Addition of increasing amounts of Sar1A (H79G) did not influence the
GST‐Sec16B (35‐194) binding (see fig 8, lanes 1‐4). As controls we monitored the
recruitment of the COPII subunit Sec23. Sec23 responded in a dose‐dependent manner to
Sar1A (H79G) activation (see fig 8, lanes 2‐4 and 11). The inhibited form of Sar1A (T39N) did
not recruit Sec23, as previously shown (see fig 8, lanes 5‐7 and 12) [18‐21]. Membrane‐
bound GST‐Sec16B (35‐194) did not affect Sec23 recruitment by Sar1A (H79G) (see fig 8,
lanes 2‐4). These results are in agreement with the hypothesized role of Sec16 as an early
initiator of ERES assembly.

146
Figure 8 ‐ Sec16B (35‐194) associates
with ER microsomes independently
of Sar1A activation ‐ ER microsomes
were derived from NRK cells, and
incubated at 32°C with RLC, and an
active form of Sar1A (Sar1A (H79G))
to promote COPII recruitment or an
inactive form of Sar1A (Sar1A (T39N))
that inhibits COPII recruitment. 1 μg
GST‐Sec16B (35‐194) fragment was
added to the reaction to examine the
fragments dependence of Sar1A
activation for recruitment. Recruited
membranes were collected by
centrifugation and examined by western blotting. Sec23 was monitored as control of microsome quality and activity by the
ability to recruit COPII using a rabbit antibody raised in‐house against a GST‐tagged full‐length Sec23 purified from E.Coli . As
this antibody also recognized the GST‐tag of Sec16B (35‐194), it was also used to detect the GST‐Sec16B (35‐195) on the same
blot. Lane 1 clearly shows that the fragment associates with the ER microsomes even in the absence of Sar1A. Although a
minor fraction precipitates out of the reaction independently of the addition of ER microsomes‐ as can be seen in lane 8 ‐ a
robust microsome dependent recruitment is still seen in the form of greater band intensity (lanes 1‐7, 9 & 10). Adding
increasing amounts of Sar1A (H79G) (0.1, 0.5 & 1 μg) does not alter the band intensities, indicating that GST‐Sec16B (35‐194)
is recruited to ER microsome independently of Sar1A activation (lanes 2‐4). Adding increasing amounts of Sar1 (T39N) neither
changes nor alters the recruitment efficiency of GST‐Sec16B (34‐195) (lanes 5‐6), indicating that the fragment does not
respond to general Sar1A activity. Sec23 was recruited from the Rat Liver Cytosol efficiently in response to Sar1A (H79G) and
was not dependent on the Sec16B fragment to be present (lane11). Sec23 is also recruited in a Sar1 (H79G) dose‐dependent
manner, as was observed in lanes 2‐4. Comparing lane 4 with lane 11 shows that COPII recruitment does not appear to be
influenced by the addition of the GST‐Sec16B (34‐195) as Sec23 band intensities remain comparably equal.
GST‐Sec16B (35‐194) maintains association with protease‐treated ER fractions of purified
Rat Liver Microsomes (ER‐RLM).
We were next prompted to look at the mechanism for GST‐Sec16B (35‐194) membrane
binding, as neither Sar1A activation nor COPII assembly influenced membrane binding.
Binding may be mediated by interactions with lipids or with peripherally associated or
integral membrane proteins. To examine these different possibilities, ER microsomes were
generated from Daugley Sprague rat livers (ER‐RLM) using established fractionation
protocols [22, 23]. The membranes were washed with increasing concentrations of KCl or
with 2.5 M urea to remove peripherally associated proteins. To examine the contribution of
proteins to the observed binding, ER‐RLM's were further treated with different proteases
for 40 min on ice to digest membrane‐associated proteins (see fig 9A). The proteolytic
activity was verified by following the cleavage of the cytosolic domain of an ER‐membrane
protein Sec12 as previously reported [18‐20, 24]. Proteolysis markedly reduced recruitment
of Sec23/24 to membranes possibly reporting on the loss of stabilizing interactions with
cargo proteins and proteins such as Sec16 itself (see fig 9A, lanes 4‐14). The binding of GST‐
Sec16B (35‐194) to microsome membranes was neither affected by salt washes nor

147
protease treatment, indicating that either binding to a membrane‐bound protease resistant
protein or direct lipid recognition mediates membrane binding for this fragment (see fig 9A
and B).
Trypsin proteolysis was also verified by Western blotting with an antibody raised specifically against Sec12, a known
membrane‐bound protein associated with ER membranes (see B). GST‐Sec16B (35‐194) recruitment was maintained both
after α‐Chymotrypsin digestion (lanes 13 & 14 (results from a parallel experiment)), and after Thermolysin digestion (lanes 11
& 12) implying, that the fragment may bind to ER membranes either through a protease resistant membrane‐bound protein
or through non‐protein interactions – possibly lipid associations. B) Trypsin control digestion of Sec12 on ER‐RLM – ER‐RLM
were either washed with 2.5 M urea or digested with trypsin for 40 min on ice and Sec12 was examined after recruitment
assays. Sec12 did not lose membrane association when washed with urea, whereas trypsin digestion caused an expected
band shift [17]. Sec12 did not respond to added Sar1A (H79G) as previously reported [11‐13].
EGFP‐Sec16B (35‐194) is not targeted to ERES in transiently transfected HeLa cells
Given the ability of the proposed Sec16B targeting signal to bind membranes, we analyzed
the ability of the domain to assemble on and mark ERES as previously reported [13]. Sec16B
(35‐194) was cloned into a mammalian expression vector with an EGFP tag and expressed
transiently in HeLa cells for 24 h. Samples were grown on coverslides, fixed and stained
against markers for different compartments of the biosynthetic transport pathway. EGFP‐
Sec16B (35‐195) was found to localize throughout cells (see fig. 10).
Figure 9 ‐ A) Protease treatment does not
inhibit GST‐Sec16B (35‐194) association with
ER‐RLM ‐ ER fractionated Rat Liver
Microsomes were generated and membrane
proteins were digested with either a buffer
control, 100 μg/mL trypsin, 5 mg/ml α‐
chymotrypsin or 5 mg/mL thermolysin for 40
min on ice. GST‐Sec16B (35‐194) recruitment
was examined as described in Fig. 6 and
efficiency of proteolysis was monitored by
blotting against Sec12 in the case of trypsin
(B), or monitoring reduced efficiency of Sec23
recruitment. Microsome dependent
recruitment could be seen as intensification
of the GST‐Sec16B (35‐194) specific band
when compared to controls without addition
of ER‐RLM (lanes 1 & 2). Lanes 3‐6 show
microsome activity prior to protease
digestion, notice background Sec23
recruitment activity when adding RLC (lane
5), which is markedly amplified with the
addition of Sar1A (H79G). Trypsin‐treated
membranes (lanes 7 & 8) still maintain robust
GST‐Sec16B (35‐194) recruitment capability
whereas COPII recruitment is markedly
effected as can be seen by the decrease in
band intensity for recruited Sec23.

Figure 10 ‐ Localization of transient EGFP‐Sec16B (35‐194) expression ‐ Transient EGFP‐Sec16B (35‐194) expression in
HeLa maintained at 37°C , fixed and co‐stained against Sec13. First image shows the fragment localizing to the cytosol
and the nucleus. The cytosolic fraction in the first image shows distinct reticular patterning in agreement with the
findings that Sec16B associates with ER membranes. No ERES specific co‐localization with Sec13 could be observed
(image 2‐4). Furthermore, EGFP‐Sec16B (35‐194) does not influence the distribution of Sec13‐marked ERES.
148
Yet it showed some reticular patterning in agreement with ER membrane binding. In some
cases the fragment was also found within the nucleus, which was interpreted as an artifact
arising from the ability of EGFP to target the nucleus. The fragment never assembled in
defined sites or punctae that could be co‐localized with ERES/COPII markers, but largely
maintained a uniform reticular expression pattern. These results suggest that the domain
contains membrane‐binding capabilities, but lacks ERES targeting properties (see fig 10).
EGFP‐Sec16B (35‐235) does not target ERES in transiently transfected HeLa cells
We further attempted to reproduce the Bhattacharyya D. & Glick B.S. observations where
the Sec16B fragment comprising of residues 35‐235 targeted ERES [13]. Sec16B (35‐235)
was isolated and cloned into a mammalian expression vector. The point mutation at
position 195 was reverted by replacing glutamine 195 with arginine and verified by
sequencing to match the fragment published by Bhattacharyya D. & Glick B.S. The fragment
was expressed uniformly throughout the cytosol of the cell, with some reticular patterning,
indicating association with ER. The fragment was never found in punctae that co‐localized
with markers for ERES/COPII. The same was found to be the case when we attempted to re‐
produce the findings of Bhattacharyya D. & Glick B.S. following the provided protocol of
their initial report (see fig. 11) [13].

Figure 11‐ EGFP‐Sec16B (35‐235) expression and localization ‐ EGFP‐Sec16B (35‐235) was expressed in HeLa cells
maintained at 37°C, fixed and co‐stained against Sec31A. No Sec31A ERES‐specific co‐localization can be observed, as
the fragment expresses uniformly throughout the cell contrary to the observations of Bhattacharyya D. & Glick B.S. [6].
Furthermore, EGFP‐Sec16B (35‐235) does not influence the distribution of Sec31A‐marked ERES, in agreement with the
observations of Budnik A. et al [18].
149
It is not clear why the results obtained by Bhattacharyya D. & Glick B.S. could not be
reproduced, however and importantly, our observations were verified by Budnik A. et al.
[25] , who cloned and expressed the same fragment and also did not detect ERES
localization. Budnik A. et al. also noted that the Sec16B (35‐235) fragment – and various
truncations of this region – were indeed expressed uniformly throughout the cell, in
agreement with the observations reported here. Overall, our analysis supports a model in
which specific targeting of Sec16B to ERES likely resides within the CCD – as recently
proposed [17]. The ability to associate with lipids likely resides in the upstream N‐terminal
region, and is, presumably, controlled by the CCD to support ERES targeting [17].
GST‐Sec16A (1096‐1190) associates with NRK microsomes independently of Sar1A activation
and COPII recruitment
A Sec16A fragment comprised of residues 924‐1227 was reported by Bhattacharyya D. &
Glick B.S. to be sufficient for targeting of Sec16A to ERES [13]. Additional observations by
Ivan V. et al. reported that tER binding of Drosophila Sec16 was also dependent on a tandem
stretch of arginine‐rich sequences upstream of the CCD [14]. Furthermore, ERES binding was
found to be independent of the CCD according to Bhattacharyya D. & Glick B.S. [13]. Based
upon these oservation, we hypothesized that a similar arginine‐rich stretch in the
mammalian homologue might confer or assist in ERES targeting. Sequence homology
analysis suggested that the Drosophila domain is homologous to residues 1111‐1169 in the
mammalian KIAA00310 clone [14]. To ensure that no targeting‐specific sequence would be
omitted, thereby producing an inactive truncation, we constructed a fragment that

150
extended from residue 1096 to 1190. Residues 1169‐1190 contain several arginine residues,
and it was reasoned that these were likely part of the mammalian arginine‐rich stretch and
needed to be included.
As with Sec16B, the Sec16A (1096‐1190) domain was produced and purified for in vitro
analysis using recruitment assays as described previously. The Sec16A (1069‐1190) domain
bound ER microsomes, and binding was not regulated by Sar1A activation or COPII assembly
(see fig 12A).
Further analysis of binding showed, as with GST‐Sec16B (35‐194), that binding of this
charged fragment to membranes appeared to be protein‐independent as 2.5 M urea‐
washed and/or 100 μg/mL trypsin‐treated NRK microsomes effectively recruited the
fragment (see fig 12B).
Figure 12 – A) GST‐Sec 16A (1096‐1190)
associates with ER microsomes
independently of Sar1A activation‐ ER
microsomes were derived from NRK cells
and used in recruitment assays with 1 μg
GST‐Sec16A (1096‐1190) as previously
described. Sec23 was monitored as
control. The GST‐Sec16A (1096‐1190)
showed association with NRK
microsomes independently of Sar1A
(H79G) (lanes 1‐5). Sec23 was recruited
in a dose‐dependent manner irrespective
of the presence of GST‐Sec16A (1096‐
1190) fragment.
B) Urea wash or trypsin treatment of
NRK membranes does not inhibit GST‐
Sec16A (1096‐1190) association ‐ NRK
microsomes were either washed with 2.5
M urea to remove associated proteins or
digested with 100 μg/mL trypsin to
remove membrane‐bound proteins, or
both washed and digested with urea and
trypsin. GST‐Sec16A (1096‐1190)
maintained membrane association
regardless of treatment as can be seen in
lanes 2‐5. The Sec23 control recruitment
shows reduced activity in response to
Sar1A (H79G) activation on the trypsin‐
digested microsomes lanes (8 & 9).
EGFP‐Sec16A (1096‐1190) does not target ERES in transiently transfected HeLa cells
The 1096‐1190 fragment of Sec16A was next examined for ERES targeting in transient
transfections of HeLa cells using a mammalian expression vector containing the EGFP‐

151
tagged fragment. As with the Sec16B fragments, Sec16A (1096‐1190) was found to localize
uniformly throughout the cell with a discernible reticular patterning, but also with strong
localization in the nucleus (see fig 13). The fragment did not assemble into defined sites or
punctae in the cell, and did not co‐localize with ERES/COPII markers (data not shown).
EGFP‐Sec16A (924‐1227) targets to the nucleus
As the Sec16A (1096‐1190) fragment did not exhibit ERES targeting, we further re‐examined
the targeting of the Sec16A (924‐1227) domain that was previously shown by Bhattacharyya
D. & Glick B.S. to be sufficient for ERES targeting [13].
Figure 13 ‐ Transient EGFP‐Sec16A (1096‐1190) expression ‐ Transient
EGFP‐Sec16A (1096‐1190) expression in HeLa cells maintained at 37°C,
collected and fixed. EGFP‐Sec16A (1096‐1190) expressed uniformly
throughout the cell with a reticular patterning when observed in the
cytosol.
The region was amplified by PCR, cloned into an EGFP expressing mammalian vector and
verified by sequencing. Transient transfection into HeLa cells showed the fragment targeting
strongly to the nucleus, frequently aggregating within this organelle (see fig 14). Some
minor punctate staining was also observed in the cytosol, with no defined association at
ERES (see fig 14, arrows). Recent analyses by Hughes H. et al. have now localized the
targeting activity of Sec16A to a larger fragment in agreement with our analysis [17].
Figure 14 ‐ Transient EGFP‐Sec16A (924‐1227) expression ‐ Transient
EGFP‐Sec16A (924‐1227) expression in HeLa cells maintained at 37°C,
collected and fixed. EGFP‐Sec16A (924‐1227) targets the nucleus with
minor punctate staining visible in the cytosol (arrows).

Cell Culture
152
Materials and Methods
HeLa were maintained at sub‐confluence in Dulbecco's Modified Eagle's Media (DMEM) (HyClone Fisher‐Scientific) supplemented with up
to 10 % Fetal Bovine Serum (Serum Source International, Inc.) and 5 % Penicillin‐Streptomycine (Cellgro) under standard incubation
conditions(37°C, 5 % CO2).
All cell lines were washed in modified DPBS without Calcium and Magnesium (HyClone Thermo Scientific) before passage using Trypsin‐
EDTA (Cellgro) to release surface adherence and diluted in their preferred culture medium.
Antibodies
The following antibodies were used in these experiments:
Mouse monoclonal against Sec31A (612350, BD Transduction Laboratories).
Mouse monoclonal against ERGIC53 (G1/93)(ALX‐804‐602, Enzo Life Science).
All Golgi specific antibodies were kindly provided by Dr. Adam Linstedt (Department of Biological Sciences, Carnegie Mellon University,
Pittsburgh, PA, USA).
Rabbit polyclonal against Sec16A clone KIAA00310 was kindly provided by Dr. Mitsuo Tagaya (School of Life Sciences, Tokyo University of
Pharmacy and Life Sciences, Tokyo).
Rabbit polyclonal against GST‐conjugated full length Human Sec23 was raised in house by Dr. Meir Aridor.
Rabbit polyclonal against a His‐tagged truncated Sec12 Δ390‐417 where the membrane associating C‐terminus was removed to establish a
soluble protein [26].
Secondary antibodies: All fluorophore‐conjugated antibodies were Alexa Goat anti‐ mouse or rabbit (Invitrogen).
Transfection
DNA transfections were performed with Effectene Transfection Reagent (Qiagen) according to provided protocol, but with optimized DNA
amounts to ensure optimal protein expression.
Cloning
pGEX‐4T‐1‐Sec16B (35‐194) was constructed by 2‐step PCR, inserting a stop codon at position 195 using Cloned Pfu‐Polymerase AD
(Stratagene) and the following primers:
Sec16s (R/Q195Stop) F: 5'‐GCTTCCAACTCTGGATAGGAGTGGCCGGGGGAG‐3'
Sec16s (R/Q195Stop) R: 5'‐CTCCCCCGGCCACTCCTATCCAGAGTTGGAAGC‐3'
The above stop codon insertion was performed on a previously constructed pGEX‐4T‐1‐Sec16B (35‐248), where the Sec16B fragment had
been amplified out of Sec16B (named Sec16s) clone provided by Kazusa DNA Research Institute (2‐6‐7 Kazusa‐kamatari, Kisarazu, Chiba
292‐0818 JAPAN). The Sec16B (35‐248) fragment was amplified out using Taq‐polymerase (GeneScript Corp) according to provided
protocol, adding a BamH I site at the 5'‐end and an Xho I site at the 3' end with the following primers:
Sec16s BamH I aa:35 F: 5'‐ GAGAGATGGATCCCATCGGCCTGTCCCTCACTCTTGGC‐3'
Sec16S234Xho‐rev: 5'‐GTCAGTACATCAGAGATGCCCCGGAGCGGGTAACTCGAGAA‐3'
pEGFP‐Sec16B (35‐194) was constructed by PCR amplification of the fragment with Taq Polymerase according to provided protocol. The
fragment was amplified from the Kazusa clone adding a BamH I site at 5' end using the above mentioned Sec16s BamH I aa:35 F primer,
and adding a Stop codon followed by a Hind III site at the 3' end with the below provided primer. Both restriction sites were used to clone
the fragment into pEGFP‐C1:
Sec16s aa:195Stop Hind III R: 5'‐GGAAACAGCTCCCCCGGAAGCTTTCATCCAGAGTTGG‐3'
pEGFP‐Sec16B (35‐234 (R195Q)) was constructed by PCR amplification of the fragment from the Kazusa clone adding a BamH I site at 5'
end using the above mentioned Sec16s BamH I aa:35 F primer, and adding a Stop codon followed by a Xba I site at the 3' end using the
below mentioned primer. Both restriction sites were used to clone the fragment into pEGFP‐C1:
Sec16s(S235Stop) Xba I R: 5'‐GCATCTCTGATGTACTGACTGAGTCTAGATTAGCTGGAGCTGAGACCAGACTC‐3'
Reversion of the arginine at position 195 to Glutamine was done by 2‐Step PCR using Cloned Pfu‐Polymerase AD and the following
primers:
Sec16s (A1187G)(Q195R ) F: 5'‐GCTTCCAACTCTGGACGGGAGTGGCCGGGGGAGC‐3'
Sec16s (A1187G)(Q195R ) R: 5'‐GCTCCCCCGGCCACTCCCGTCCAGAGTTGGAAGC‐3'
pGEX‐4T‐1‐Sec16A (1096‐1190) and pEGFP‐Sec16A (1096‐1190) were constructed by PCR amplification of the fragment from the Kazusa
clone (KIAA00310) using Taq‐ Polymerase, adding a BgI II site at the 5' end and a BamH I site at the 3' end (see the primers below). The
fragment was cloned into a BamH I site of either pGEX‐4T‐1 or pEGFP‐C1:
Bgl II Sec16L aa: 1096 F: 5'‐GGAGATCCAGGTAGATCTGATCGTTACC‐3'

153
Sec16L aa: 1190 Bam HI R: 5'‐CGAGTGGGAGCTGGATCCGCTGCGGCGG‐3'
Orientation was verified by PCR using Taq‐Polymerase and the following primers:
pGEX‐4T‐1 (841‐856) F: 5'‐CCAGCAAGTATATAGC‐3'
EGFP Insert Seq II F: 5'‐CCAACGAGAAGCGCG‐3'
Sec 16L ABS PCR Check R: 5'‐CCGGGGATCCGCTGCGG‐3'
pEGFP‐Sec16A (924‐1227) was constructed by PCR amplification of the fragment from the from the Kazusa clone (KIAA00310) adding a Bgl
III site at 5' and adding a Stop codon followed by a Xba I site at the 3' end using the following primers. Both inserted restriction sites were
used to clone the fragment into a BamH I site in pEGFP‐C1:
Sec16L Bgl II (aa:924) F: 5'‐GCCCAGAACTCAGCACAGTCAAGATCTAGTCTGGTTCTGGTCGACGCGGG‐3'
Sec16L aa:1227Stop Bam HI R: 5'‐CCACTGCTGAAATTGCTGCGGGATCCTTAGTAGGCAAAATCGCCG‐3'
Orientation was verified by PCR using Taq‐Polymerase and the above mentioned primers.
All constructs were additionally verified by sequencing carried out by Genewiz, Inc. pGEX‐4T‐1 were verified using primers T7 and T7 Term
provided by Genewiz (see below):
T7: 5'‐TAA TAC GAC TCA CTA TAG GG‐3'
T7 Term: 5'‐GCT AGT TAT TGC TCA GCG G‐3'
Whereas pEGFP‐C1 clones were verified by the above mentioned EGFP Insert Seq II F, and the EGFP Insert Seq Prim R constructed primer
(see below):
EGFP Insert Seq Prim R: 5'‐CCATTATAAGCTGCAATAAACAAG‐3'
All primers were acquired from Integrated DNA Technologies, Inc. (IDT)
Temperature Block Assay
HeLa cells were transfected as described and incubated at 37°C for 14‐16 h. Media was supplemented with 20 mM HEPES (pH=7.4) (Fisher
Scientific) and incubated at 15°C or 10°C for 4 h on an aluminum block ¾ submerged in a closed water bath placed in a 4°C cold room.
Samples were recovered and fixed in 3.7 % formaldehyde (Sigma‐Aldrich) solution in modified DPBS without Calcium and Magnesium
(HyClone Thermo Scientific).
Immunofluorescence
General Immunofluorescence: Cells were seeded onto 12 mm circular glass cover slides (Fischer Scientific) at a density optimized to reach
app. 80 % confluence at time of fixation. Slides recovered at their defined time points/stages were all fixed in 3.7 % Formaldehyde Solution
(Sigma‐Aldrich) in modified DPBS without Calcium and Magnesium (HyClone Thermo Scientific), incubated 20 min. at RT, then washed 3
times with modified DPBS without Calcium and Magnesium (HyClone Thermo Scientific). Slides were usually stored at 4°C prior to staining.
Slides were washed 3 time with 0.05 % Saponin (Sigma‐Aldrich) in modified DPBS without Calcium and Magnesium (HyClone Thermo
Scientific) (0,05 % Saponin‐PBS Solution) to ensure proper plasma membrane permeabilization before blocking with 5 % Goat Serum
(Sigma‐Aldrich) in 0.05 % Saponin‐PBS Solution, incubation for 20 min. at RT. All primary antibodies were diluted to optimized relations in
0.05 % Saponin‐PBS and left on the cover slides for 45‐60 min incubations at RT. Cover slides were washed 3 times with 0.05 % Saponin‐
PBS before adding 1 to 500 dilutions of secondary fluorophore‐conjugated antibody solutions, incubate 15 min. at RT, washed twice with
0.05 % Saponin‐PBS and once with regular modified DPBS without Calcium and Magnesium (HyClone Thermo Scientific) before mounting
on precleaned Superfrost® Microscope Slides (12‐550‐143, Fisher Scientific) with Fluoromount G (Electron Microscopy Sciences) and left to
air‐dry O.N. before sealing with clear nail polish.
Slides were visualized on an Olympus Fluoview 1000 using a PLAPON 60 x objective with a NA = 1.42. Images were processed using the
provided software (FV10‐ASW version 02.00.03.10 (Olympus Corporation) and Adobe Photoshop CS3 (Adobe Photoshop Version: 10.0.1
(Adobe)).
Protein Purification
Expression vectors were transformed into E. Coli BL 21 (Invitrogen) and grown up to an OD≈0.6 at 37°C. Protein production was induced by
adding up to 0.1 mM isopropyl‐β‐D‐thiogalactoside (IPTG) (FisherBiotech) and incubation at 37°C for an additional 4h.
Bacteria were pelleted down and usually stored at ‐80°C until further processing.
For His‐tagged purification, pellets were re‐suspended in 50/100/1 TNE (50 mM Tris‐HCl (pH=8.0) (EMD), 100 mM NaCl (EMD) and 1 mM
EDTA (Sigma‐Aldrich)) supplemented with 1 mM PMSF (Sigma‐Aldrich), and further supplemented with 1 mM GDP (Sigma‐Aldrich) and 10
mM β‐Mercaptoethanol (J.T. Baker).
For GST‐tagged purification pellets were re‐suspended in 50/10/1 TNE (50 mM Tris‐HCl (pH=8.0)(EMD), 10 mM NaCl (EMD) and 1 mM
EDTA (Sigma‐Aldrich)) supplemented with 1 mM PMSF (Sigma‐Aldrich).
For both types of purification 46,900 u/mL Lysozyme from Chicken egg white (Sigma‐Aldrich) was added and the re‐suspension was
allowed to incubate at 4°C for 30 min before complete lysis of cells was achieved by 3 cycles of freeze/thawing between N2 (l) and a 32°C
water bath.
Up to 10 mM MgCl2 (Fisher Scientific) and 1,700 u/mL DNase I (Roche) was added, lysates were incubated an additional 30 min at 4°C to
remove genomic DNA.
Cell debris was removed by centrifugation at 22000 x g for 30 min.

154
GST‐tagged protein purification: Protein was bound upon GST‐Sepharose 4B (GE Healthcare Life Science), incubation 1‐1 ½ h incubation at
4°C. Beads were washed thoroughly with PBS over several rounds before Elution in 50/150 TN (50 mM Tris‐HCl (pH=8.7)(EMD), 150 mM
NaCl (EMD)) supplemented with 15 mM reduced Glutathion (Sigma‐Aldrich). Bound protein was eluted over 4 rounds of 3 mL Elution
Buffer.
High yield elutions were pooled and reduced to a volume below 3 mL with Macrosep 10K Omega Centrifugal Device (Pall Life Science)
before dialysis into 25 mM HEPES (pH=7.4)(Calbiochem), 125 mM KOAc (Fisher Scientific) using 7000 MWCO Slide‐A‐Lyzer Dialysis Cassette
(Thermo‐Scientific). Final concentrate was alliquoted, flash frozen in N2(l) and stored at ‐ 80°C.
His‐Tagged protein purification: Protein was bound to Ni‐NTA Agarose (Qiagen) and washed once with
50/100/1 TNE, and then with 40 mL of 50/100/1 TNE supplemented up to 0.3 M NaCl (EMD) and 1 mM MgCl2 (Fisher Scientific). The
beads were further washed in 40 mL 50/300/1 HNE Buffer (50 mM HEPES (pH=7.4)(Calbiochem), 300 mM NaCl (EMD), 50 μM EGTA (Fisher
Scientific), 1 mM MgCl2 and 10 mM β‐Mercaptoethanol (J.T. Baker), and an additional 40 mL of 25 mM Imidazole (EMD) in 50/300/1 HNE
Buffer adjusted to pH=7.4 before elution in 2 mL fractions with 500 mM Imidazole (EMD) in 50/300/1 HNE Buffer adjusted to pH=7.4.
Protein rich elutions were pooled and concentrated using Macrosep 10K Omega Centrifugal Device (Pall Life Science) before dialysis into
25 mM HEPES (pH=7.4)(Calbiochem), 125 mM KOAc (Fisher Scientific) using 7000 MWCO Slide‐A‐Lyzer Dialysis Cassette (Thermo‐
Scientific). Final concentrate was alliquoted, flash frozen in N2(l) and stored at ‐ 80°C.
Microsome Recruitment Assay
NRK derived Microsomes and Rat Liver Cytosol were prepared according to Plutner H. et al [27]. Rat liver Derived ER specific microsomes
(ER‐RLM) were prepared using an adapted discontinuous gradient fractionation protocol according to Balch, W. et al [23], modified for
preparation from Rat liver . Briefly, crude homogenate from the Livers of 2 female Daugley Sprague rats (10 mM Tris‐HCL (pH=7.4)(EMD), 5
mM EDTA (Sigma‐Aldrich), 1 mM PMSF (Sigma‐Aldrich), 0.1 TIU/mL aprotinin (Sigma‐Aldrich), 5 μg/mL leupeptin (Sigma‐Aldrich), was
centrifuged at 3000 rpm (JA‐20 Beckman), supernatant and "off‐white" upper pellet was collected and adjusted to 1.3 M Sucrose
(FisherBiotech)(12 mL), placed on top of 2.4 M sucrose bottom (2 mL) anf overlaid with 1.2 M sucrose and 0.8 M sucrose (14 mL and 7 mL
respectively) in a total volume of 35 mL. After centrifugation at 9000 x g the membrane fraction at interface 1.2 M and 1.3 M sucrose was
collected as a fraction enriched in ER microsomes. The fraction was adjusted to 0.4 M sucrose and collected by centrifugation (100000 x g
for 1 H at 4 °C). The fraction was tested for enrichment of Sec12 and ability to recruit Sec23. Recruitment assays were modified and carried
out according to Aridor, M. et al 1995 [18] Aridor, M. et al 1998 [19] and Aridor, M. & Balch, W 2000 [20]. Using 20‐40 μg of membranes
that were either un‐washed, salt washed with 0.5‐2 M KCL (EMD) or 2.5 M urea (Merck) , or treated with either 100 μg/mL Trypsin (Sigma‐
Aldrich), 5 mg/ml α‐Chymotrypsin (Sigma‐Aldrich) or 5 mg/mL Thermolysin (Sigma‐Aldrich) for 40 min. on ice. Microsomes were
incubated in 35 mM HEPES (pH=7.4)(Calbiochem), 2.5 mM MgOAc (J.T. Baker), 80 mM KOAc (Fisher Scientific), 5 mM EGTA (Fisher
Scientific), 0.2 mM GTP (Sigma‐Aldrich), 1 mM ATP (Sigma‐Aldrich), 5 mM Creatine Phosphate (Sigma‐Aldrich), 0.2 u of rabbit muscle
creatine phosphokinase (Sigma‐Aldrich), supplemented with either GST‐Sec16B (35‐195), GST‐Sec16A (1096‐1190) or/and His‐Sar1 (H79G)
or His‐Sar1 (T39N) in the amounts mentioned for each assay in a final volume of 60 μL. Reaction was carried out of 32°C in a water bath
for 15 min followed by further 10 min of incubation on ice. Membranes were collected by centrifugation through a sucrose cushion.
Samples were added on top of 180 μL 15 % sucrose, 75 mM KOAc (Fisher Scientific) and 2 mM MgOAc (J.T. Baker), and spun down at
16000 x g for 15 min at 4 °C, liquid was aspirated, and samples re‐spun at 16000 x g for an additional 5 min to remove excess liquid.
Samples were re‐suspended in 2 x Sample Buffer.
SDS‐PAGE
Gels and electrophoretic separation were performed according to standard protocol [28].
Western Blot
SDS‐PAGE gels were wet transferred to Protran® Nitrocellulose Transfer Membrane (Whatman®, Schleicher & Schuell) in a Mini Trans‐Blot
Electrophoretic Transfer Cell (Bio‐Rad), under optimized transfer conditions specific for the Polyacrylamide content, in 20 % methanol
(EMD), 25 mM Tris‐HCL (EMD) and 200 mM Glycine (Bio‐Rad). Membranes were blocked for 1 h in 5 % Non‐Fat Instant Dry Milk solution of
1 % Tween‐20‐ TBS. Membranes were incubated 14‐16 h at 8°C with optimal dilutions of primary antibodies in 5 % Non‐Fat Instant Dry
Milk solution of 1 % Tween‐20‐ TBS. After 3 times of 10 min washes in 1 % Tween‐20‐ TBS, membranes were developed using HRP
conjugated Goat antibodies (Thermo Scientific) targeted towards the species of the primary antibody diluted 1 to 5000 in 5 % Non‐Fat
Instant Dry Milk solution of 1 % Tween‐20‐ TBS. After 45 min. incubation at RT and 3 times of 10 min washes in 1 % Tween‐20‐ TBS, the
blots were recorded on HyBlot CL X‐Ray film (Denville Scientific Inc.) using either SuperSignal West Dura Extended Duration Substrate
(Thermo Scientific) or HyGLO (Denville Scientific Inc.) according to provided protocols.

155
References
1. Ong, Y.S., et al., p125A exists as part of the mammalian Sec13/Sec31 COPII subcomplex to
facilitate ER‐Golgi transport. J Cell Biol, 2010. 190(3): p. 331‐45.
2. Mizoguchi, T., et al., Determination of functional regions of p125, a novel mammalian
Sec23p‐interacting protein. Biochem Biophys Res Commun, 2000. 279(1): p. 144‐9.
3. Aravind, L., The WWE domain: a common interaction module in protein ubiquitination and
ADP ribosylation. Trends in biochemical sciences, 2001. 26(5): p. 273‐5.
4. Zweifel, M.E., D.J. Leahy, and D. Barrick, Structure and Notch receptor binding of the tandem
WWE domain of Deltex. Structure, 2005. 13(11): p. 1599‐611.
5. Inoue, H., et al., Roles of SAM and DDHD domains in mammalian intracellular phospholipase
A1 KIAA0725p. Biochim Biophys Acta, 2012. 1823(4): p. 930‐9.
6. Ferro‐Novick, S., et al., Yeast secretory mutants that block the formation of active cell surface
enzymes. J Cell Biol, 1984. 98(1): p. 35‐43.
7. Novick, P., C. Field, and R. Schekman, Identification of 23 complementation groups required
for post‐translational events in the yeast secretory pathway. Cell, 1980. 21(1): p. 205‐15.
8. Schekman, R., et al., Yeast secretory mutants: isolation and characterization. Methods
Enzymol, 1983. 96: p. 802‐15.
9. Shaywitz, D.A., et al., COPII subunit interactions in the assembly of the vesicle coat. J Biol
Chem, 1997. 272(41): p. 25413‐6.
10. Espenshade, P., et al., Yeast SEC16 gene encodes a multidomain vesicle coat protein that
interacts with Sec23p. J Cell Biol, 1995. 131(2): p. 311‐24.
11. Gimeno, R.E., P. Espenshade, and C.A. Kaiser, COPII coat subunit interactions: Sec24p and
Sec23p bind to adjacent regions of Sec16p. Mol Biol Cell, 1996. 7(11): p. 1815‐23.
12. Yorimitsu, T. and K. Sato, Insights into structural and regulatory roles of Sec16 in COPII
vesicle formation at ER exit sites. Molecular biology of the cell, 2012.
13. Bhattacharyya, D. and B.S. Glick, Two mammalian Sec16 homologues have nonredundant
functions in endoplasmic reticulum (ER) export and transitional ER organization. Mol Biol
Cell, 2007. 18(3): p. 839‐49.
14. Ivan, V., et al., Drosophila Sec16 mediates the biogenesis of tER sites upstream of Sar1
through an arginine‐rich motif. Mol Biol Cell, 2008. 19(10): p. 4352‐65.
15. Saraste, J. and E. Kuismanen, Pre‐ and post‐Golgi vacuoles operate in the transport of Semliki
Forest virus membrane glycoproteins to the cell surface. Cell, 1984. 38(2): p. 535‐49.
16. Mezzacasa, A. and A. Helenius, The transitional ER defines a boundary for quality control in
the secretion of tsO45 VSV glycoprotein. Traffic, 2002. 3(11): p. 833‐49.
17. Hughes, H., et al., Organisation of human ER‐exit sites: requirements for the localisation of
Sec16 to transitional ER. J Cell Sci, 2009. 122(Pt 16): p. 2924‐34.
18. Aridor, M., et al., Sequential coupling between COPII and COPI vesicle coats in endoplasmic
reticulum to Golgi transport. J Cell Biol, 1995. 131(4): p. 875‐93.
19. Aridor, M., et al., Cargo selection by the COPII budding machinery during export from the ER.
J Cell Biol, 1998. 141(1): p. 61‐70.
20. Aridor, M. and W.E. Balch, Kinase signaling initiates coat complex II (COPII) recruitment and
export from the mammalian endoplasmic reticulum. J Biol Chem, 2000. 275(46): p. 35673‐6.
21. Kuge, O., et al., Sar1 promotes vesicle budding from the endoplasmic reticulum but not Golgi
compartments. The Journal of cell biology, 1994. 125(1): p. 51‐65.
22. Balch, W.E., et al., Reconstitution of the transport of protein between successive
compartments of the Golgi measured by the coupled incorporation of N‐acetylglucosamine.
Cell, 1984. 39(2 Pt 1): p. 405‐16.
23. Balch, W.E., B.S. Glick, and J.E. Rothman, Sequential intermediates in the pathway of
intercompartmental transport in a cell‐free system. Cell, 1984. 39(3 Pt 2): p. 525‐36.

156
24. Nakano, A., D. Brada, and R. Schekman, A membrane glycoprotein, Sec12p, required for
protein transport from the endoplasmic reticulum to the Golgi apparatus in yeast. J Cell Biol,
1988. 107(3): p. 851‐63.
25. Budnik, A., K.J. Heesom, and D.J. Stephens, Characterization of human Sec16B: indications of
specialized, non‐redundant functions. Scientific reports, 2011. 1: p. 77.
26. Weissman, J.T., H. Plutner, and W.E. Balch, The mammalian guanine nucleotide exchange
factor mSec12 is essential for activation of the Sar1 GTPase directing endoplasmic reticulum
export. Traffic, 2001. 2(7): p. 465‐75.
27. Helen Plutner, C.G., Xiaodong Wang, Paul LaPointe, and William E. Balch, Microsome‐Based
Assay for Analysis of Endoplasmic Reticulum to Golgi Transport in Mammalian Cells. 1 ed.
Cell Biology Assays: Essential Methods, ed. F.J.C.E. Geri Kreitzer. Vol. 1. 2010: Academic
Press: Elsevier Inc.
28. Gallagher, S.R., One‐dimensional SDS gel electrophoresis of proteins. Curr Protoc Mol Biol,
2006. Chapter 10: p. Unit 10 2A.

157
Conclusions, Discussion and
Perspectives
Summary of findings
In this thesis, we have investigated the COPII accessory protein p125A and the COPII
scaffolding protein Sec16. The overall aim of this project was to identify mechanisms that
respond to changes in the local lipid environment, and the effect of these mechanisms on
the overall assembly of ERES and progression of ER export. A brief overview of the findings
are presented below. These will be discussed in more detail in the following sections.
We show that membrane binding and ERES targeting of p125A is controlled by its DDHD
domain. The DDHD domain recognizes and binds to monophosphorylated
phosphatidylinositols, especially to PI(4)P and to some degree also PA and PS. Inhibition of
the DDHD lipid binding domain causes reduced membrane association and ERES localization
of p125A, as seen for the (850KGRKR/EGEEE854)(PI‐X) and the ΔDDHD mutants, which
abrogate specific lipid recognition. The effects of the abrogated lipid recognition are further
explored using kinetic analysis (FRAP). Wild type p125A promotes an apparent retention of
the COPII coat at ERES. This retention is perturbed when measuring the kinetics of p125A
(PI‐X), which shows faster kinetic rates at ERES, implying a destabilization of the interaction
of p125A with membranes.
We demonstrate that the lipid recognition of the DDHD domain is influenced by a novel
mechanism. Oligomeric interactions between p125A SAM domains can control the
specificity of the DDHD domain's intrinsic lipid recognition. We show that inhibition of SAM‐
mediated oligomerization, by introducing an interaction targeted mutation (L690E) at the
assembly interface, causes p125A to lose ERES targeting and membrane binding.
Furthermore, interference of p125A membrane binding leads to a dominant negative effect
and disrupts ERES. This loss of targeting and the effects on ERES can be partially rescued by
substituting the DDHD with the PI(4)P targeting PH domain of Fapp1.

158
We demonstrate that p125A associates and segregates with ER exit sites when incubated at
low temperatures, under conditions that induce inhibition of cargo transport at either the
ERGIC or ERES.
Expanding upon the temperature‐blocking experiments, we find that ERES marked by
p125A, Sec31A and Sec23 separate from Sec16A and Sec16B at the low temperature
conditions. Sec16A and Sec16B are instead collected in large structures, which separate
from ERGIC53, Golgi, and ERES. These observations suggest that Sec16A and Sec16B act at
the ERES prior to the assembly of the COPII cage.
To explore further on these findings, we examine the correlation between p125A and
Sec16A during high levels of overexpression of p125A and mutants thereof. The
overexpression of p125A wt leads to the formation of large aggregate sites. Presumably,
some of the structures represent coated enlarged ERES. These sites are largely segregated
from Sec16A. In contrast, during overexpression of a p125A mutant where the lipid‐binding
domain has been deleted and where its ability to oligomerize is inhibited, the aggregates
formed become engulfed by Sec16A. These observations imply that p125A promotes the
displacement of the COPII coat from the Sec16A scaffolding. These findings furthermore
suggest that p125A provides linkage between the two COPII layers (Sec23/24‐Sec13/31)
during the assembly of vesicles at the ERES. The displacement and linkage is dependent on
p125A recognizing specific lipid signals at ERES – likely PI(4)P.
The consequences of the perturbed p125A lipid recognition is additionally examined in
relation to ER export. We confirm that depletion of p125A causes perturbation of the steady
state transport levels in cells as reported by the dominant dispersion of the Golgi [1]. This
dispersion can be rescued by re‐introducing an RNAi‐resistant clone of p125A. In contrast,
this dispersion cannot be rescued by introducing a dominant negative RNAi‐resistant clone
where both the mutation in the SAM domain (L690E) together with the abrogation of the
lipid binding in the DDHD domain (PI‐X) have been introduced.
These observations lead us to propose the following model: Sec16A associated with both
the inner and outer layer of the COPII coat, provides initial scaffolding at initiating ERES.
Recruitment of p125A to the Sec16A scaffold occurs as part of the recruitment of
Sec13/Sec31 since p125A exists as an integral component associated with the outer layer

159
[1]. p125A oligomerization via SAM domains and changes in the ERES local lipid
environment due to induced recruitment of kinases, i.e. PI4KinIIIα, promote the p125A
DDHD domain to bind to PI(4)P [2, 3]. The lipid binding, in turn, promotes p125A‐controlled
linkage between the two COPII layers and furthers the displacement of Sec16A from the
budding vesicle site. The stabilization of the coat is now instead provided by p125A and the
displacement of Sec16A directs further progression and maturation of the forming vesicle.
In an additional analysis we identify a unique WWE binding motif that is part of the p125A
P‐Q rich domain [4, 5]. We hypothesize that this motif is responsible for p125A association
with Sec31A.
Finally, we attempt to verify previously identified regions in Sec16A and Sec16B responsible
for ERES targeting. We expand upon these regions membrane binding capabilities, and find
that these regions do confer ER membrane binding. But these regions do not cause specific
targeting of the proteins towards ERES.
The findings in this project provide a better understanding of the molecular mechanism for
regulatory control of the COPII machinery at the vesicle bud site.
SAM – a domain for oligomerization
SAM domains consist of a 70‐residue stretch that forms a compact 5‐helix bundle with a
globular fold. The helix bundle consists of 4 short helices (α1‐α4) and long C‐terminal helix
(α5). The 4 short helices interact with the N‐terminal part of α5 to form a characteristic
hydrophobic pocket [6‐9]. The roles of the SAM domains are numerous and diverse. SAM
domains are mostly found in context of larger multi‐domain proteins located in all cellular
compartments, reflecting their participation in a wide variety of different processes [10].
The capability to modulate function by homo‐ or hetero‐oligomerization is a general
characteristic of SAM domains. Several versions of SAM domains have been shown capable
of polymerizing into larger functional structures [6‐9]. Potential lipid binding capabilities by
SAM domains have also been reported [11, 12]. Associations between SAM domains have
been mapped to two patches of residues, an apolar mid‐loop (ML) surface patch near the
center of the peptide, and an end‐helix (EH) surface patch where the important residues are
provided by α5 [8].

Prior to this work, the SAM domain in p125A was not functionally characterized. A novel
160
screen developed to identify SAM domains with polymerization capabilities identified p125A
SAM as having borderline oligomeric behavior, but this finding was not further explored
[13].
The diacylglycerol kinase (DGK) δ SAM domain polymerizes into larger sheet like structures
in the presence of Zn 2+ [14]. Likewise, the highly homologous p125A SAM oligomerized and
precipitated in response to in vitro exposure to Zn 2+ . Polymerization was highly dependent
on an EH leucine residue at position 690 within the α5 helix. The physiological relevance of
p125 SAM assembly was demonstrated by the impact of the p125A L690E mutation on ERES
stability based on morphological and kinetic readouts. Although lipid recognition has
previously been reported for some domain family members, the p125A SAM domain by
itself did not exhibit any lipid recognition activity or membrane binding, as measured by a
lipid overlay assay and cellular localization analysis. Similarly, the double mutant of the full‐
length p125A, containing both the L690E and the DDHD PI‐X mutations, was predominantly
cytosolic with minimal ERES targeting.
Mapping possible interaction partners of p125A SAM is of significant interest, as the
regulatory functions of p125A SAM are not completely understood. Is the regulatory
function the sole consequence of homotypic p125A SAM interactions, or do other factors
with homologous SAM domains participate in forming a larger regulatory unit? A potential
co‐player could be DAGKδ. The SAM‐possessing DAGKδ is localized to ER membranes, and
ERES localization could very well involve associations with p125A. ERES targeting is
abrogated by deletion of the DAGKδ‐SAM domain, and the domain is required for DAGKδ‐
mediated regulation of ER export [15]. Our study focused mainly on homotypic interactions
of p125A. Utilizing the purified p125A SAM domain in pull‐down experiments followed by
mass spectrometry analysis ought to determine whether the domain interacts with other
components – such as DAGKδ, or even the smaller p125B homolog – to regulate ER export.
It would also be of interest to create a “super active” SAM domain in p125A to monitor ERES
organization. Substituting p125A SAM with a SAM domain known to form tighter
associations, e.g. the SAM domain of the transcription factor translocation ETS leukemia
(TEL) might be one possibility [8]. Since TEL SAM may only recognize itself, it would be

161
interesting to observe whether re‐targeting of p125A towards the nucleus may occur. The
introduction of a stronger SAM‐SAM chimeric protein interaction may on the other hand
displace and perturb the functions of endogenous p125A instead. Alternatively, a simple
tandem duplication of the domain may accomplish the desired effect. Such chimeras would
be useful to study the connection between SAM domains and lipid recognition. They would
also provide a tool to examine how the lipid composition influences budding and homotypic
fusion after Sar1 GTP hydrolysis. A super‐active p125A should exhibit extended ERES
binding. This binding would address the role of the COPII cage in recruiting proteins that
promote vesicle fission. Prolonged or enhanced association would also be useful to identify
additional factors necessary for establishing an efficient cargo transport after budding.
An important question that warrants further investigation is the potential physiological role
of Zn 2+ in the oligomerization reaction. Zn 2+ ‐mediated SAM oligomerization was identified in
the synaptic scaffolding Shank proteins, suggesting a role in synaptic plasticity. Upon Zn 2+ ‐
binding, Shank‐SAM domains form dense sheets composed of helical fibers. Similar
structures were observed in cryo‐EM images underneath the postsynaptic membranes of
synapses. It was hypothesized that local influx of Zn 2+ leads to denser packing of Shank
proteins and increased proximity between Shank associated factors, thus promoting the
probability of enzymes binding to their respective substrates [16, 17]. Similarly, local
elevations of Zn 2+ at ERES may promote p125A associations with itself and with COPII
components, and thus promote retention of COPII cage ERES.
p125A SAM may have further functionalities beyond oligomerization. At present no definite
screen exists to elucidate other potential functions.
DDHD domains and the influence of lipid recognition
DDHD domains are mainly defined by sequence homology. It is speculated that DDHD
domains may bind metal ions and that the domain may be utilized for selective lipid binding
[18‐20]. Only one study has so far tried to define the biochemical properties of the DDHD
domain in the cell [21]. Membrane targeting has been identified for all currently known
members of the DDHD family [22‐24]. Our work provides the first functional
characterization of the DDHD domain in p125A. We demonstrate that the p125A‐DDHD is
involved in selective lipid binding. Furthermore, we find that this lipid binding is controlled

162
by adjacent regulatory elements – p125A (SAM) ‐ which in the case of p125A promote
assembly.
Moreover, we define a short amino acid stretch within the domain (850‐KGRKR‐854)
required for lipid recognition. Reversing the charge on this stretch (EGEEE – PI‐X) causes loss
of apparent lipid‐binding specificity. This leads to a significant increase in exchange rates of
the mutant p125A at ERES. We assume that these changes target the actual lipid binding
site of the DDHD domain, since the introduced mutations improved our purification yield of
the p125A fragment (643‐989). The improved yield is an indication that the domain does not
missfold or aggregate. Furthermore, full length p125A (PI‐X) maintains solubility in vivo and
does not aggregate when transiently expressed thus further supporting proper folding.
p125A (PI‐X) maintains its ability to associate with its protein binding partners. A similar
phenotype is also observed when the DDHD is deleted. Thus, we conclude that the lipid‐
binding site of p125A was targeted.
The smaller p125A homolog, p125B, is a PLA1 that hydrolyzes primarily PA and to a lesser
degree PS and PC. It is ubiquitously expressed and is predominantly cytosolic with a limited
population bound to cis‐Golgi membranes [25]. In contrast to p125A, p125B does not
include an N‐terminus that binds to COPII. Interestingly, chimeras where the p125A N‐
terminus has been added to p125B become retargeted to ERES. Similar chimeras in which
the N‐terminus of p125A is fused to the C‐terminus of another DDHD family protein,
DDHD1, do not localize to ERES, perhaps because DDHD1 lacks a SAM domain [20, 26].
Overall, these observations imply that membrane binding by DDHD domains might be co‐
incidental binding mediated by distal motifs such as SAM domains (our work and [21]), or
FFAT motifs as found in the Nir family of DDHD containing proteins (Nir 1‐3) [24]. A potential
strategy for exploring this hypothesis would be to switch the DDHD domains from the Nir
protein with the DDHD domains of p125A, B or DDHD1. Would an obvious change in activity
be noticed for the chimeric proteins if the DDHD domain was under the control of a SAM
domain instead of an active lipase domain? Would a chimeric protein's targeting be changed
if the DDHD domain is under the influence of an FFAT motif instead of a SAM domain?
Does the p125A DDHD bind ions as proposed in Nir 1‐3 [18, 19]? It has been shown that
family member DDHD1 gets recruited to membranes in response to elevated Ca 2+ levels by

163
ionomycin treatment [20]. However, depletion of Ca 2+ by addition of the high affinity Ca 2+
chelator EGTA to budding reactions in permeabilized cells does not inhibit vesicle budding.
Ca 2+ depletion only inhibits a later fusion stage between formed VTC's and Golgi
membranes. Therefore, the COPII machinery is functional in a Ca 2+ ‐depleted environment
[27]. Furthermore, studies of the regulator of COPII, ALG‐2, which binds Sec31A and controls
coat retention at ERES in response to elevated Ca 2+ levels, used p125A as a marker for ERES.
The study shows that p125A associates with ERES and does not respond to Ca 2+ depletion
[28, 29]
The defining DDHD residues of the domain are believed to be responsible for ion binding
[18, 19]. Studying the consequences of mutating these residues in vivo and in vitro would
address whether the ion‐binding motif of the DDHD domain is essential for the domain
function. This could be monitored in the full‐length protein by studying the cellular effects in
transient transfections, thus addressing whether a potential ion binding influences the
p125A ERES targeting. FRAP analysis with these constructs would further address whether
ion binding influences the binding stability and dynamics of p125A at the ERES. Influence on
lipid specificity should be examined by introducing the DDHD mutations into the p125A
(643‐989) fragment and measure the fragment's activity in lipid‐blot overlay assays.
Monitoring and adjusting local lipid distribution might be an additional function of p125A.
DDHD‐containing proteins have been identified to participate in lipid transport [24, 30].
p125B has furthermore been identified to possess lipase activity [25, 31]. Although p125A
does present a lipase domain, no such activity has yet been identified for p125A [21, 25, 32].
As the lipid binding site for the DDHD domain has not been determined, mapping of the
binding site should be performed to address whether DDHD domains and DDHD‐containing
proteins play a role in local lipid homeostasis.
WWE domain of p125A – a possible Sec31A binding motif
The WWE domain of p125A may be responsible for the binding to Sec31A through the
unstructured region within Sec31A. The unstructured region of Sec31A binds to both Sec23A
and controls the GTPase rate of Sar1 [33, 34]. Binding of p125A to this region might suggest
a molecular model of COPII regulation involving both COPII layers and p125A. p125A WWE
binding to Sec31A would cause displacement of the Sec31A catalytic tryptophan and

asparagine from the hydrolysis reaction in the GTP‐binding pocket in Sar1A. This in turn
164
would promote retention of COPII at the ERES.
Sec31p in yeast contains an unusually high amount of serine, threonine and proline residues
in its carboxy‐terminal half, which contains the unstructured region of Sec31p [35, 36].
These residues were identified as part of a PEST motif (rich in (P) proline, (E) glutamic acid,
(S) serine and (T) threonine), which is associated with protein stability and degradation [37].
Phosphorylation of PEST motifs is thought to control protein stability by regulating
recruitment of F‐box‐containing ubiquitin E3 ligases that are known to target proteins for
proteasomal degradation [35, 36]. The study found that Sec31 was phosphorylated mainly
on serine residues, and that phosphatase‐treated Sec31/13 fractions inhibited vesicle
formation in vitro [37].
WWE motifs are regularly identified as part of E3 ubiquitin ligases that are known to bind to
phosphorylated or poly(ADP ribosyl)'ated (PAR) targets [4, 5]. Interactions between p125A
WWE and Sec31A through minor or extensive post‐translational phosphorylation of Sec31A
may be a very possible regulatory mechanism. Elevated levels of phosphorylated Sec31A
may promote either higher affinity or avidity of p125A WWE binding to the Sec31A
unstructured region.
Very recent findings have suggested that Caseine Kinase 2 (CK2) phosphorylation of Sec31A
decreases the Sec31 latency time at bud sites and gives rise to a larger cytosolic population
of Sec31 [38]. The study furthermore implied that CK2 phosphorylation of serines positioned
at residues 527 and 799 in Sec31 inhibits Sec31‐Sec23 interactions. These observations
suggest that the p125A WWE more likely recognizes PAR modifications of Sec31, as our
study implies that p125A association with the COPII layers retains and maintains the COPII
cage at budding ERES.
Mutants of Sec31A harboring selective deletions within its unstructured region combined
with analysis of p125A binding using co‐IP or pull‐down assays with GST‐tagged p125A WWE
could define p125A binding sites. Site‐directed mutagenesis can be further utilized to define
whether phosphorylation of serines in Sec31A regulates binding to the WWE domain of
p125A.

165
The binding of p125A to Sec31A has been localized to the last 180 aa of Sec31A (1041‐1220)
[1]. Furthermore, Alg‐2, an identified accessory protein, influences functional ERES assembly
by binding to Sec31A in the proline‐rich unstructured region (aa 800‐1113) in response to
locally elevated levels of Ca 2+ [28]. Based upon these results, we hypothesized that p125A
WWE, similar to Alg‐2, might bind within the unstructured region of the human Sec31A. As
the Sec31A unstructured region stretches from residues 800‐1091, we estimated that the
binding site would most likely reside within a region of Sec31A comprising residues 1041‐
1091 [29, 39]. However, we were not able to detect binding between p125A and a GST‐
Sec31A (1041‐1091) fragment in pull‐down assays. We were also not able to identify any
association between an EGFP‐tagged Sec31A (1041‐1091) fragment with endogenous or
Flag‐tagged p125A in co‐IP experiments (data not shown). The lack of interaction may be
ascribed to lack of post‐translational modifications (e.g. phosphorylation, PAR or even
ubiquitination [37, 40]) of Sec31A that may take place during membrane binding. Indeed,
transiently expressed EGFP‐Sec31A (1040‐1091) in HeLa cell was predominantly cytosolic,
suggesting that the fragment is not targeted to ER membranes (data not shown).
Another useful option for examining p125A influence on Sec31A would be to create a hybrid
Sec31A protein, substituting the structural elements of the human Sec31A with homologous
structural elements from S. cerevisae Sec31p. These elements are structurally conserved
and would likely exhibit behavior homologous to the human version when expressed with
the human unstructured region and vice versa [41, 42]. Since S. cerevisiae does not express
a protein homologous to p125A, it is expected that there would be no binding between the
hybrid protein expressing the yeast unstructured loop and p125A. It would furthermore be
interesting to monitor whether the hybrid Sec31 exhibits different regulation. This would
provide a tool to examine p125A‐independent Sec31A regulation. This approach could
identify novel accessory proteins that bind Sec31A and are involved in COPII regulation.
Alternatively, p125A WWE binding might also occur within the C‐terminal structured α‐
solenoid region of Sec31A. α‐solenoid structures are involved in protein‐protein
interactions. For example, ankyrin repeats that fold in an α‐helix‐turn‐α‐helix structure that
is typical for α‐solenoids are known to be involved in protein‐protein interactions during
signal transduction, as observed for the Notch receptor [5, 43‐45]. Purification and pull‐
down experiments of GST‐tagged fragments from within the Sec31A C‐terminal α‐solenoid

166
could be one approach to address whether the Sec31A C‐terminal α‐solenoid is a binding
site for p125A WWE. This approach should be additionally verified through co‐IP of epitope
tagged versions of the same fragments after transient expression in mammalian cells.
Whether p125A associates with any part of the unstructured region of Sec31A should be
addressed.
Mutagenesis of the WWE domain aimed at inhibiting p125A‐Sec31A binding is also of
interest. If successful, such an approach may unravel the role of p125A in linking the inner
and the outer layers of the coat and how p125A regulates coat configuration. Such
experiments may provide tools to address the role of inner and outer layer linkage in cargo
packaging for efficient ER export. Regulation of coat configuration may be of particular
interest when studying packaging of larger cargo. For example pro‐collagen, as perturbation
of Sec31A binding to Sec23 is known to cause defects in collagen secretion. Moreover,
p125A defects have also been suspected to cause impaired collagen secretion [46, 47].
Our findings that p125A is required to displace Sec16A from the ERES provides the first
evidence of p125A's role in promoting the progression of COPII budding and vesiculation. It
is assumed that Sec16A binding to both Sec23 and Sec31A inhibits the cage components in
linking. Inhibiting the assembly of Sec23 and Sec31 within the catalytic pocket of Sar1
stabilizes the ERES nucleation. Sec16A thereby prevents premature disassembly of the COPII
cage [34, 48, 49]. p125A displaces Sec16A in response to the locally elevated PI(4)P
concentrations at the ERES. p125A furthermore stabilizes the linkage between the two
layers through its binding with both Sec23 and Sec31. This in turn promotes the association
of Sec23 and Sec31 within the nucleotide pocket of Sar1 and as a consequence promotes
the hydrolysis of the pocket‐bound GTP.
It is not difficult to expand upon this model adding a more regulatory role of p125A during
the GTP hydrolysis in response to variations in the local lipid composition. We and other
groups have shown that the DDHD domain of p125A has the capability of recognizing
additional lipid variants beyond PI(4)P, e.g. monophosporylated PI's, PA and PS [21], and
that the specificity is modulated by SAM oligomerization. With these observations in mind,
we propose that p125A binding to charged lipids, such as PI(4)P, at ERES promotes a general
stabilization of p125A with itself and with the lipid membrane. This in turn also promotes

the previously mentioned COPII linkage and stabilization as well as the hydrolysis of the
Sar1‐bound GTP. Thus, p125A regulates and controls the COPII cage at the ERES during
167
budding and vesiculation. This model should be possible to test in conditions using artificial
liposomes with predefined compositions. Addition of Sar1, Sec23/Sec24, Sec13/31 and
p125A to liposomes with low levels of PI(4)P should exhibit a slower rate of GTP hydrolysis
when compared to liposomes composed with higher concentrations of PI(4)P. These
experiments would address the influence of PA and PS on p125A‐mediated decoding of the
lipid environment, and whether these lipids also promote COPII linkage, stabilization and
Sar1‐bound GTP hydrolysis.
Sec16A and Sec16B collect into structures at low temperature incubation
Our observations that Sec16A and Sec16B assemble into defined structures that segregate
from COPII and also ERGIC and Golgi when cargo export is blocked either at the ERGIC or at
the ERES, is surprising. The degree of segregation suggests that the main function of
mammalian Sec16 (mSec16) occurs at an early stage during ER export likely promoting ERES
nucleation and initial COPII assembly. The localization of the mSec16 at distinct sites in the
tER such as the defined cup like structures described by Hughes H. et al [50] is an additional
indication that mSec16 probably functions prior to ERES assembly [50‐52]. Controlled
release from the temperature block while monitoring mSec16, cargo and COPII behavior
would be instrumental in defining an actual role for the mSec16 structures. Temperature
block release experiments may also answer whether Sec16 binds cargo. In addition, such
experiments could also define the role of mSec16 in cargo loading, as it is known that
mSec16 levels influence ER‐to‐Golgi transport, as well as ERES distribution and size during
acute or chronic cargo load [2, 53].
It has also been reported that Sec16A and Sec16B are capable of associating with each other
both in homo‐oligomeric and hetero‐oligomeric complexes [54, 55]. The clustering at low
temperatures may very likely be driven by such oligomerization. The dot1 mutation in the
CCD of the P. pastoris Sec16p is presumed to inhibit Sec16p oligomerization, which
manifests itself as a lack of ERES organizational clustering [51]. The influence of the Sec16A
and Sec16B homo‐oligomers on the ERES organization should be explored. This could be
investigated by introduction of a homologous dot1 mutation in the CCD into the mammalian

168
Sec16A and Sec16B. If the observed clustering during low temperature incubations is
inhibited, this might imply that Sec16A and Sec16B homo‐oligomers are used to maintain a
certain degree of ERES organization. If the clustering is not observed, it would imply that
Sec16A and Sec16B associate with an ER‐bound component that remains to be resolved.
Another question that should also be addressed is, whether the homo‐oligomers have any
influence upon ERES and cargo dynamics. Deletions of the Sec16p CCD or substitution of the
Sec16p CCD with the ACE1 domain from S. cerevisiae Sec31p still renders the yeast viable
[55], implying that the CCD might be redundant in lower eukaryotes. In mammalian systems,
the CCD may promote a higher degree of ERES organization and thereby also a higher
degree of cargo packaging efficiency, which seems to be implied when ER export is observed
in Sec16A knock‐down experiments [2].
Recent observations from Montegna E.A. et al. [56] suggest that Sec16A associates with
Sec12 at tER sites. Prior observations of Sec12 at 15°C have shown that the protein
segregates from ERES at 15°C and does not leave the ER [57]. As these observations did not
utilize the 10°C block, it remains to be analyzed whether Sec12 is co‐localizing with the
mSec16 structures observed at 10°C. Furthermore, the Montegna E.A. et al. observations
implied that recruitment of mSec16 to the ERES correlates with Sec12‐Sar1 interaction [56].
It would therefore be pertinent to study if Sec16A and Sec12 co‐localize or segregate at low
temperature incubations. Such analysis would begin to address the roles of Sec12 prior to
COPII recruitment.
The most interesting observations from the temperature blocks, is the efficient reduction in
the dispersed pool of mSec16 at 10°C. Consistent with our observations, Iinuma T. et al. [32]
identified a population of Sec16A in the cytosol by subcellular fractionation. This study could
furthermore demonstrate an increase of membrane‐bound Sec16A in response to the
addition of Sar1‐GTP using microsome recruitment assays. Endogenous Sec16A has also
been demonstrated in vivo to accumulate on juxtanuclear areas during transient expression
of Sar1‐GTP[53]. The pool of cytosolic mSec16, and likely a non‐ERES associated ER
membrane‐bound pool of mSec16, are both probably maintained dynamically by p125A‐
mediated displacement of mSec16. At 15°C and 10°C the release of cargo vesicles is either
slowed down or blocked. This results in less to no p125A‐mediated displacement and
recycling of mSec16, thereby collecting the dispersed mSec16. The mSec16 is instead bound

169
at complexes formed prior to ERES nucleation from which it cannot proceed. The
composition of these complexes, besides mSec16, needs still to be examined. Temperature
block experiments will undoubtedly provide a very powerful tool in the future dissection of
Sec16A and Sec16B functions and associations at the ERES.
The membrane‐associating regions of mSec16 that have been examined seem to bind
membranes independently of specific activation such as Sar1A recruitment ([50, 52, 58] and
own observations). Furthermore, the fact that there is a cytosolic population of mSec16
[32], suggests that the ER membrane dissociation involves either masking of the membrane‐
associating regions of mSec16, or more likely modifications of Sec16 such as
phosphorylations through e.g. ERK7 (a.k.a MAPK15) [59]. These modifications may perhaps
cause repulsion of the protein from the charged membrane. Indeed, Zacharogianni M. et al.
[59] have reported that the tER targeting domain of D. melanogaster Sec16 (dSec16)
contains multiple phosphorylation sites. It would therefore be interesting to examine the
dynamics of mSec16 in a MAPK15‐depleted environment. It would also be of interest to
examine whether a significant decrease of the dispersed mSec16 and visible mSec16
structures are observed under the MAPK15 depleted conditions.
p125A mediated displacement of Sec16A from ERES
We find that overexpression of p125A mutants impaired in lipid recognition also have an
ability to collect mSec16A around aggregated ERES. In contrast, overexpression of p125A wt
shows a clear separation of p125A from mSec16A. This suggests that Sec16A dissociation
from ERES occurs in response to p125A recognizing and binding to a specific subset of lipids
at the ERES, in particular PI(4)P [2, 3]. The mechanisms of this exchange have yet not been
explored. Sec16A is known to bind to the majority of the components of the COPII
machinery [32, 49, 50, 54, 60‐65]. A possible mechanism might comprise of a disruption or
inhibition of the Sec23 and Sec31A associations with Sec16A due to the p125A mediated
Sec23‐Sec31 linkage in response to lipid binding causing [1]. The lipid binding may promote
conformational change of p125A. This conformational change brings the two coat
components closer together and thereby promotes their association/linkage, in addition to
the Sec16A displacement. This mechanism implies that Sec16A acts as a stabilizing scaffold
during the early stages of ERES formation and assembly. Furthermore, Sec16A may also be

170
responsible for providing and maintaining a minor population of COPII cage components in
response to Sec12‐Sar1A activation and membrane deformation [56, 66].
Our findings support and expand the proposed model of Sec13‐Sec31 mediated
displacement of the Sec16p‐Sec13 tetramer [55]. We propose that Sec16A association with
Sec31A inevitably also includes p125A in the formed complex during the early stages of ERES
formation [1, 63]. Sec16A promotes the associations between Sec31A and Sec23 and
thereby also controls the associations between Sec31A‐Sec23 with Sar1 [33, 34, 49, 60‐65,
67]. This may account for the observed delay in the GTP hydrolysis of Sar1 on microsomes
and at the ERES in the presence of Sec16 [49, 68]. The delay of Sar1 hydrolysis may
subsequently promote the Sar1 membrane deformation activity. Sec16A furthermore
promotes the association between p125A and Sec23. As the ERES formation progresses,
p125A responds to a local elevation of PI(4)P at the ERES by binding to the lipid and
displacing Sec31A and Sec23A from its association with Sec16A [2, 3]. This explains our
observations of Sec16A displacement or lack thereof during overexpression of either wt or
mutant p125A, respectively. This may also explain the findings of Sec16A localizing in cup‐
like structures surrounding the ERES [50]. Sec16A function might therefore actively
participate in controlling the localization of membrane bending and subsequent tubulation.
An interesting experiment would be to perform siRNA‐mediated knock‐down of PI4KinIIIα
and monitor the distribution of Sec16A in these conditions. Would Sec16A persist for longer
periods at the few ERES observed? What would be the distribution between the displaced
pool of Sec16A compared to the collected population on the ER membrane measured by
fractionation? As we assume that Sec16A displacement from ERES occurs in response to
p125A lipid binding, one would assume that Sec16A would show a similar collection as seen
during low temperature incubations and likely a pronounced co‐localization with the few
established ERES [2]. Similar observations would also be apparent in a p125A‐depleted
environment, where Sec16A would associate more readily with the ER membrane and
exhibit a more pronounced dispersion on the ER correlating with the Sec31A staining
pattern [1, 32].
Our model also gives an explanation for the dispersal of the Sec31A pattern during p125A
depletion [1, 32]. The nucleation of new ERES is initiated by Sec12 recruiting and tethering
Sar1 to the ER [57, 69‐71]. Nucleation does not occur in predefined ERES, but is rather

171
initiated in an intermediate environment between rER and sER that both promotes
tubulation and is in the vicinity of the protein production machinery. These intermediary
potential sites can be found throughout the entire ER [72, 73]. The nucleation event
subsequently recruits Sec16A, which stabilizes the Sar1‐induced membrane deformation.
Whether the coat components are recruited with Sec16A or follow after Sec16A association
remains to be determined. Our observations with Sec16A separating from Sec31A during
low temperature incubation, suggests that the outer layer of COPII is recruited at a separate
stage or as a separate event to Sec16A recruitment. Likely, the Sec13/31A/p125A complex is
recruited after Sec16A association with Sec12 and Sar1 at the nucleation site.
The association of Sec16A with Sar1 and the coat components targets PI(4)P‐Kinases to the
nucleation site [3]. The local elevation of PI(4)P triggers lipid binding of Sec31A‐bound
p125A, and in turn promotes Sec16A displacement as well as p125A‐mediated COPII linkage
and stabilization. Naturally, stable ERES will be formed more frequently in an environment
that exhibits a higher level of PI(4)P phosphorylation, i.e. the interface between ER and
Golgi.
ERES nucleation events occur throughout the ER regardless of the presence of p125A.
However, in a p125A‐depleted environment the stabilization of the newly formed pre‐ERES
is maintained by Sec16A that scaffolds the COPII at the site [56, 66]. The lack of p125A‐
mediated Sec16A displacement retains Sec31A at several potential ERES, causing a clear
dispersion of the Sec31A expression pattern [1, 32].
The interplay between Sec16A and the COPII components in relation to p125A function
needs to be further examined. The present results clearly indicate that p125A plays an
important role during the segregation of COPII from Sec16A scaffolding within the ERES.
Introducing mutations and deletions within the p125A binding domains for Sec31A and
Sec23 should be used to examine the relation between p125A‐promoted COPII linkage and
p125A‐promoted Sec16A displacement from ERES [1, 21, 74]. Would these mutations cause
collection of Sec16 on the ER membrane as seen in the temperature blocks? Or would Sec16
be less apparent at ERES and exhibit a higher level of cellular dispersion? The dynamics
should also be examined in these conditions by both FRAP and by measuring VSV‐G

172
transport. This will establish the influence of the Sec16A interplay with p125A lipid binding
during ERES formation and cargo loading.
Abrogating the associations between Sec16A and Sec23 as well as Sec31A should be
explored by modifications of their respective Sec16A binding domains [49, 60‐65]. This
would provide a better time line of the recruitment events occurring during COPII‐
dependent ERES assembly. It would further define the scaffolding role of Sec16A during ER
export. Additionally, this can be used to distinguish the influence of p125A linkage and
stability of the COPII cage after Sec16A displacement [1, 21, 74]. What would be the
consequences for ERES formation if Sec16 cannot interact with Sec23 or Sec31A? If visible
ERES are formed, what is the turnover of Sec16A, Sec23, Sec31A and p125A measured by
FRAP during these conditions?
An interesting paradox is that Sec16p is not required for budding in experimental settings
with artificial liposomes [48]. However, Sec16p is still a vital component of the secretory
pathway and for survival in yeast [61]. It would be interesting to examine, whether the
scaffolding activity of mammalian Sec16 might be dispensable for in vivo budding by
establishing a knock‐out animal or cell line. RNA interference experiments already indicate
that Sec16 may have a predominantly supportive role in the cell, since traffic is merely
delayed under these conditions [2].
Does the displacement of Sec16A occur in response to PI(4)P? Or do some of the other
acidic lipids, i.e. PA or PS, promote Sec16A displacement? These questions should be
answered by monitoring the dynamics of ERES nucleation. In particular, the dynamics of the
p125A‐mediated displacement of Sec16A in response to changes in the local acidic lipid
environment should be tested. An experimental setup with artificial liposomes has already
been provided Supek F. et al. [48]. The influence of mSec16A, in correlation to p125A wt as
well as the single and double mutants, on the budding activity from synthetic liposomes
should be measured. These experiments should be compared to the earlier described
examinations of p125A‐lipid binding and COPII linkage on synthetic liposomes in this
discussion. This would address the influence of mSec16A scaffolding on ERES nucleation and
COPII‐mediated budding. This would also aid in showing whether p125A‐mediated
displacement of mSec16A does promote a higher level of budding activity.

173
Sec16A and Sec16B membrane binding and ERES targeting
Clearly, ERES targeting to and participation in ERES assembly is not encompassed in the 35‐
194 region of Sec16B nor the 924‐1227 region of Sec16A, as previously suggested [54, 58].
However, our analysis suggests that these regions may provide binding avidity to ER
membranes to support ERES localization. Salt washes and proteolytic analysis suggest that
lipid recognition may be the likely mode for the domains to associate with the membranes
at the tER sites. Although protease‐resistant membrane‐bound protein cannot be rule out as
a possible binding partner. Further investigations of the mSec16 membrane binding regions
are required to define their specificity. Possible strategies would be to introduce various
truncations or mutations in the arginine‐rich region preceding the CCD, which is known to
be essential for ERES targeting [52, 75]. This would add additional knowledge to the
targeting of each Sec16 protein. This would likely also aid in answering whether the
targeting of mSec16 is dependent on specific lipids, or whether they bind to a specific
membrane protein.
The required residues for membrane binding are hard to map: for one, no apparent cationic
or hydrophobic stretches are easily identified in either protein. Secondly, structural and
sequence alignment tools have not been able to identify homology between the Sec16
proteins and known lipid or protein‐binding sequences or regions. Thus, detailed structure‐
function analysis of the CCD should be performed. Introducing minor deletions or sequence
variations and testing them in similar recruitment assays as described in the chapter:
"Investigations in p125A‐Sec31A associations and mammalian Sec16A and B membrane
binding", may be one method of dissecting the functionality of these regions.
It is pertinent to verify that Sec16 association is lipid dependent and not protein associated.
An obvious method would be to examine the recruitment of both the Sec16B (35‐194)
fragment and the Sec16A (1096‐1190) fragment to various synthetic liposomes of varying
phospholipid composition without the supplement of cytosol. This will circumvent adding
any membrane association factor. Moreover, this approach would also provide a potential
platform for dissecting which lipids might be mediating the binding. It would furthermore
identify, whether the fragments bind by specific lipid recognition or by less specific
hydrophobic interactions. An approach could be to establish an RNAi screen and select for

174
candidates with membrane tethering features that produce Sec16 phenotypes similar to
those seen at the 10°C incubation. Identified candidates would subsequently be verified for
co‐localization by microscopy using the 10°C ER exit block, and further tested for protease
resistance by microsome digestion assays as described in the previous chapter.
Sec16 has been shown to be recruited to ER microsomes in a Sar1A‐dependent manner [32].
Functions of Sec16 at the ERES likely involve regulated associations with the COPII in a
sequential fashion prior to the vesicle formation, as implied in our presented paper. Sec16
association with ERES appears to be transient, as shown by the temperature blocks. The
question arises whether Sec16 delays the assembly of the COPII coat at newly forming ERES.
Alternatively, the association of Sec16 with COPII plays a role in establishing a coat‐enriched
platform that initiates the assembly of the tER, as suggested from studies in P. pastoris [51].
Recent findings in S. cerevisiae identified a Sec16p fragment comprising the residues 565–
1235 that inhibited Sar1 hydrolysis [68]. Thus, Sec16 seems to have a role in both tER
assembly and coat retention.
Physiological relevance of p125A regulation
Even though a p125A knock‐out mouse has been reported [76], a complete characterization
of the phenotype is yet to be published. Since p125A is not essential during development
[77] and the mice are viable, the role of p125A during development remains to be defined.
Since the protein is ubiquitously expressed throughout the organism, it is reasonable to
assume it must partake in general COPII regulation. This implies that there is a functional
redundancy with other COPII regulators, or that cargo transport can tolerate deregulated
conditions derived from p125A ablation during embryogenesis. A potential candidate for
functional redundancy might be p125B [25, 31, 78]. To address whether p125B confers
redundancy, p125B should be depleted in the background of p125A depletion. The analysis
will not only provide insight in the roles of this smaller homolog, but also on the global need
for the p125 family in transport regulation.
p125A‐mediated COPII regulation may play a selective role during the packaging of specific
cargo such as collagen. In a p125A‐deficient environment, collagen may be transported at a
significantly lower rate than normal conditions as observed in cells expressing mutated
Sec23A [79, 80]. Absence of p125A does cause defects in spermiogenesis in mice and in

175
particular has inhibitory effects on the formation of acrosomes [76] Furthermore, we show
that Golgi disperses upon knock‐down of p125A, in accordance with previous findings [1,
26]. This indicates that steady‐state levels of cargo transport are being affected. These
observations therefore provide evidence for a more extensive traffic inhibition that not only
affects collagen export.
What then causes the inhibition in acrosome formation? The acrosome is derived from the
Golgi [81]. Since siRNA‐mediated depletion of p125A causes inhibition of the ER export and
concomitant disassembly of the Golgi, it is evident that the knock‐out animals must display
similar defects that cause a disassembled Golgi. The p125A deficiency likely inhibits
trafficking of essential components including lipids, proteases and golgins that are important
in the formation of both Golgi and the acrosome. Studies examining p125A and COPII
controlled traffic will give a greater insight in male fertility and potentially lead to novel
targets for development of male contraceptives.
We show that overexpression of p125A leads to the formation of enlarged p125A‐coated
bud sites from which apparent budding of smaller vesicles can be identified.
Morphologically reminiscent large COPII‐coated membrane structures of 200‐500 nm in size
are also formed in response to the expression of the BTB adaptor KLHL12. KLHL12 interacts
with the N‐terminus of Sec31A and when in complex with the ubiquitin ligase CUL3, KLHL12‐
CUL3 catalyzes monoubiquitination of Sec31A. This activity is required for the secretion of
collagen [40]. COPII vesicles larger than 60‐80 nm have not been described previously and
these findings imply that the formation of vesicles accommodating large proteins such as
pro‐collagen fibers is controlled by monoubiquitination of Sec31A.
The KLHL12 driven enlargement of COPII‐coated membranes led us to examine the enlarged
p125A structures seen during overexpression experiments more thoroughly by super
resolution SIM microscopy and 3D reconstruction. We found that the p125A‐promoted
structures are complex and appear to resemble continuous membrane surfaces forming
several perforated spherical or rhomboid domains. Medium‐sized structures were rarely
compact spheres and were frequently perforated by hollow tubular structures. We were not
able to determine the morphology of the smaller p125A‐containing structures due to
resolution limits. Sec31A and Sec23 were associated with these large structures, which often

176
consisted of a continuous membrane. Overexpression of p125A and the resulting retention
of COPII coat might have caused inhibition of vesicle fission leading to the formation of
enlarged structures. p125A at these structures remained dynamic (as measured by FRAP),
suggesting that the structures may be functional intermediates in ER exit. We monitored the
movement of cargo in the form of VSV‐G through these structures. The endo H assays used
to monitor this movement show that VSV‐G export is inhibited when p125A is
overexpressed. We furthermore observed possible budding of small vesicles occurring from
these structures, which would explain the apparent maintenance of ER export.
A similar study should be conducted with KLHL12‐CUL3‐induced structures to determine
whether they maintain dynamics and ER export. These studies should furthermore also
determine whether p125A is a component of the KLHL12‐CUL3‐induced vesicles. It would
also be interesting to measure the kinetics of the KLHL12‐CUL3‐induced Sec31A structures
to determine the dynamics within these larger vesicles. Finally, SIM‐3D reconstructions of
the KLHL12‐CUL3 structures to our p125A reconstructions should be compared. Jin L. et al.
did provide EM images of KLHL12‐CUL3 induced single vesicles [40]. However, these images
do not address whether the KLHL12‐CUL3 vesicles contain cargo or whether they have
dynamic budding occurring on their surface [40]. These structures may indeed turn out to
be equivalent to the dynamic budding structures that we observe during p125A
overexpression.
Concluding remarks
The basic COPII machinery has been well characterized. Elucidation of the regulatory
mechanisms that control the organization and function of the COPII machinery at ERES is
only beginning. Our results and other recent findings are beginning to uncover a network of
COPII‐interacting proteins that are involved in ERES regulation. These regulators include Alg‐
2, Sed4, STAM's and the membrane protein TANGO1. These regulators respond to increased
cargo load or to changes in the local environment. All these proteins may utilize defined
mechanisms to control the stability and organization of the COPII coat in order to support
selective traffic activities [28, 29, 82‐84].
p125A regulates COPII in response to the local lipid environment. This provides a specific
molecular mechanism for COPII regulation that directly couples ER export to lipid signaling

177
and possibly also lipid biogenesis. This is consistent with previous reports that have
observed elevation of PI(4)P and PA levels at the ERES [3], and experiments where
PI(4)KinIIIα depletion was shown to cause dispersion of ERES [2].
As more factors are being categorized within the ER‐to‐Golgi transport, our knowledge of
how this transport influences cellular functions is becoming more expanded and complex. It
is therefore pertinent to map the basic mechanisms associated with the assembly and
regulation of the ERES and cargo loading. This will promote better understanding of events
in the early ER export stages that may lead to targeting of the formed vesicles, e.g. whether
specific cargo packaging or specific SNARE recruitment is decisive for the fate of the formed
vesicle.
This project has provided a new mechanism in the COPII‐dependent transport that connects
the functions of Sec16 with the lipid recognizing activity of p125A during the ERES
formation.

178
References
1. Ong, Y.S., et al., p125A exists as part of the mammalian Sec13/Sec31 COPII subcomplex to
facilitate ER‐Golgi transport. J Cell Biol, 2010. 190(3): p. 331‐45.
2. Farhan, H., et al., Adaptation of endoplasmic reticulum exit sites to acute and chronic
increases in cargo load. EMBO J, 2008. 27(15): p. 2043‐54.
3. Blumental‐Perry, A., et al., Phosphatidylinositol 4‐phosphate formation at ER exit sites
regulates ER export. Developmental cell, 2006. 11(5): p. 671‐82.
4. Aravind, L., The WWE domain: a common interaction module in protein ubiquitination and
ADP ribosylation. Trends in biochemical sciences, 2001. 26(5): p. 273‐5.
5. Zweifel, M.E., D.J. Leahy, and D. Barrick, Structure and Notch receptor binding of the tandem
WWE domain of Deltex. Structure, 2005. 13(11): p. 1599‐611.
6. Stapleton, D., et al., The crystal structure of an Eph receptor SAM domain reveals a
mechanism for modular dimerization. Nature structural biology, 1999. 6(1): p. 44‐9.
7. Thanos, C.D., K.E. Goodwill, and J.U. Bowie, Oligomeric structure of the human EphB2
receptor SAM domain. Science, 1999. 283(5403): p. 833‐6.
8. Kim, C.A., et al., Polymerization of the SAM domain of TEL in leukemogenesis and
transcriptional repression. The EMBO journal, 2001. 20(15): p. 4173‐82.
9. Harada, B.T., et al., Regulation of enzyme localization by polymerization: polymer formation
by the SAM domain of diacylglycerol kinase delta1. Structure, 2008. 16(3): p. 380‐7.
10. Qiao, F. and J.U. Bowie, The many faces of SAM. Science's STKE : signal transduction
knowledge environment, 2005. 2005(286): p. re7.
11. Aviv, T., et al., The RNA‐binding SAM domain of Smaug defines a new family of post‐
transcriptional regulators. Nature structural biology, 2003. 10(8): p. 614‐21.
12. Bhunia, A., et al., NMR structural studies of the Ste11 SAM domain in the dodecyl
phosphocholine micelle. Proteins, 2009. 74(2): p. 328‐43.
13. Knight, M.J., et al., A human sterile alpha motif domain polymerizome. Protein science : a
publication of the Protein Society, 2011. 20(10): p. 1697‐706.
14. Knight, M.J., et al., Zinc binding drives sheet formation by the SAM domain of diacylglycerol
kinase delta. Biochemistry, 2010. 49(44): p. 9667‐76.
15. Nagaya, H., et al., Diacylglycerol kinase delta suppresses ER‐to‐Golgi traffic via its SAM and
PH domains. Molecular biology of the cell, 2002. 13(1): p. 302‐16.
16. Baron, M.K., et al., An architectural framework that may lie at the core of the postsynaptic
density. Science, 2006. 311(5760): p. 531‐5.
17. Gundelfinger, E.D., et al., A role for zinc in postsynaptic density asSAMbly and plasticity?
Trends in biochemical sciences, 2006. 31(7): p. 366‐73.
18. Lev, S., et al., Identification of a novel family of targets of PYK2 related to Drosophila retinal
degeneration B (rdgB) protein. Mol Cell Biol, 1999. 19(3): p. 2278‐88.
19. Lev, S., The role of the Nir/rdgB protein family in membrane trafficking and cytoskeleton
remodeling. Exp Cell Res, 2004. 297(1): p. 1‐10.
20. Yamashita, A., et al., Generation of lysophosphatidylinositol by DDHD domain containing 1
(DDHD1): Possible involvement of phospholipase D/phosphatidic acid in the activation of
DDHD1. Biochim Biophys Acta, 2010. 1801(7): p. 711‐20.
21. Inoue, H., et al., Roles of SAM and DDHD domains in mammalian intracellular phospholipase
A1 KIAA0725p. Biochim Biophys Acta, 2012. 1823(4): p. 930‐9.
22. Lev, S., Non‐vesicular lipid transport by lipid‐transfer proteins and beyond. Nature reviews.
Molecular cell biology, 2010. 11(10): p. 739‐50.
23. Aikawa, Y., et al., Involvement of PITPnm, a mammalian homologue of Drosophila rdgB, in
phosphoinositide synthesis on Golgi membranes. The Journal of biological chemistry, 1999.
274(29): p. 20569‐77.

179
24. Amarilio, R., et al., Differential regulation of endoplasmic reticulum structure through VAP‐
Nir protein interaction. The Journal of biological chemistry, 2005. 280(7): p. 5934‐44.
25. Nakajima, K., et al., A novel phospholipase A1 with sequence homology to a mammalian
Sec23p‐interacting protein, p125. J Biol Chem, 2002. 277(13): p. 11329‐35.
26. Shimoi, W., et al., p125 is localized in endoplasmic reticulum exit sites and involved in their
organization. J Biol Chem, 2005. 280(11): p. 10141‐8.
27. Aridor, M., et al., Sequential coupling between COPII and COPI vesicle coats in endoplasmic
reticulum to Golgi transport. J Cell Biol, 1995. 131(4): p. 875‐93.
28. Yamasaki, A., et al., The Ca2+‐binding protein ALG‐2 is recruited to endoplasmic reticulum
exit sites by Sec31A and stabilizes the localization of Sec31A. Mol Biol Cell, 2006. 17(11): p.
4876‐87.
29. Shibata, H., et al., The ALG‐2 binding site in Sec31A influences the retention kinetics of
Sec31A at the endoplasmic reticulum exit sites as revealed by live‐cell time‐lapse imaging.
Biosci Biotechnol Biochem, 2010. 74(9): p. 1819‐26.
30. Vihtelic, T.S., et al., Localization of Drosophila retinal degeneration B, a membrane‐
associated phosphatidylinositol transfer protein. The Journal of cell biology, 1993. 122(5): p.
1013‐22.
31. Sato, S., et al., Golgi‐localized KIAA0725p regulates membrane trafficking from the Golgi
apparatus to the plasma membrane in mammalian cells. FEBS Lett, 2010. 584(21): p. 4389‐
95.
32. Iinuma, T., et al., Mammalian Sec16/p250 plays a role in membrane traffic from the
endoplasmic reticulum. J Biol Chem, 2007. 282(24): p. 17632‐9.
33. Antonny, B., et al., Dynamics of the COPII coat with GTP and stable analogues. Nat Cell Biol,
2001. 3(6): p. 531‐7.
34. Bi, X., J.D. Mancias, and J. Goldberg, Insights into COPII coat nucleation from the structure of
Sec23.Sar1 complexed with the active fragment of Sec31. Developmental cell, 2007. 13(5): p.
635‐45.
35. Rechsteiner, M. and S.W. Rogers, PEST sequences and regulation by proteolysis. Trends in
biochemical sciences, 1996. 21(7): p. 267‐71.
36. Yada, M., et al., Phosphorylation‐dependent degradation of c‐Myc is mediated by the F‐box
protein Fbw7. The EMBO journal, 2004. 23(10): p. 2116‐25.
37. Salama, N.R., J.S. Chuang, and R.W. Schekman, Sec31 encodes an essential component of the
COPII coat required for transport vesicle budding from the endoplasmic reticulum. Mol Biol
Cell, 1997. 8(2): p. 205‐17.
38. Koreishi, M., et al., CK2 Phosphorylates Sec31 and Regulates ER‐To‐Golgi Trafficking. PLoS
One, 2013. 8(1): p. e54382.
39. Tang, B.L., et al., Mammalian homologues of yeast sec31p. An ubiquitously expressed form is
localized to endoplasmic reticulum (ER) exit sites and is essential for ER‐Golgi transport. J Biol
Chem, 2000. 275(18): p. 13597‐604.
40. Jin, L., et al., Ubiquitin‐dependent regulation of COPII coat size and function. Nature, 2012.
482(7386): p. 495‐500.
41. Stagg, S.M., et al., Structure of the Sec13/31 COPII coat cage. Nature, 2006. 439(7073): p.
234‐8.
42. Fath, S., et al., Structure and organization of coat proteins in the COPII cage. Cell, 2007.
129(7): p. 1325‐36.
43. Li, J., A. Mahajan, and M.D. Tsai, Ankyrin repeat: a unique motif mediating protein‐protein
interactions. Biochemistry, 2006. 45(51): p. 15168‐78.
44. Kajava, A.V., Review: proteins with repeated sequence‐‐structural prediction and modeling.
Journal of structural biology, 2001. 134(2‐3): p. 132‐44.
45. Kobe, B. and A.V. Kajava, When protein folding is simplified to protein coiling: the continuum
of solenoid protein structures. Trends in biochemical sciences, 2000. 25(10): p. 509‐15.

180
46. McGary, K.L., et al., Systematic discovery of nonobvious human disease models through
orthologous phenotypes. Proceedings of the National Academy of Sciences of the United
States of America, 2010. 107(14): p. 6544‐9.
47. Alam, I., et al., Differentially expressed genes strongly correlated with femur strength in rats.
Genomics, 2009. 94(4): p. 257‐62.
48. Supek, F., et al., Sec16p potentiates the action of COPII proteins to bud transport vesicles. J
Cell Biol, 2002. 158(6): p. 1029‐38.
49. Yorimitsu, T. and K. Sato, Insights into structural and regulatory roles of Sec16 in COPII
vesicle formation at ER exit sites. Molecular biology of the cell, 2012.
50. Hughes, H., et al., Organisation of human ER‐exit sites: requirements for the localisation of
Sec16 to transitional ER. J Cell Sci, 2009. 122(Pt 16): p. 2924‐34.
51. Connerly, P.L., et al., Sec16 is a determinant of transitional ER organization. Curr Biol, 2005.
15(16): p. 1439‐47.
52. Ivan, V., et al., Drosophila Sec16 mediates the biogenesis of tER sites upstream of Sar1
through an arginine‐rich motif. Mol Biol Cell, 2008. 19(10): p. 4352‐65.
53. Watson, P., et al., Sec16 defines endoplasmic reticulum exit sites and is required for secretory
cargo export in mammalian cells. Traffic, 2006. 7(12): p. 1678‐87.
54. Bhattacharyya, D. and B.S. Glick, Two mammalian Sec16 homologues have nonredundant
functions in endoplasmic reticulum (ER) export and transitional ER organization. Mol Biol
Cell, 2007. 18(3): p. 839‐49.
55. Whittle, J.R. and T.U. Schwartz, Structure of the Sec13‐Sec16 edge element, a template for
assembly of the COPII vesicle coat. J Cell Biol. 190(3): p. 347‐61.
56. Montegna, E.A., et al., Sec12 Binds to Sec16 at Transitional ER Sites. PLoS One, 2012. 7(2): p.
e31156.
57. Weissman, J.T., H. Plutner, and W.E. Balch, The mammalian guanine nucleotide exchange
factor mSec12 is essential for activation of the Sar1 GTPase directing endoplasmic reticulum
export. Traffic, 2001. 2(7): p. 465‐75.
58. Hughes, H. and D.J. Stephens, Sec16A defines the site for vesicle budding from the
endoplasmic reticulum on exit from mitosis. J Cell Sci, 2010. 123(Pt 23): p. 4032‐8.
59. Zacharogianni, M., et al., ERK7 is a negative regulator of protein secretion in response to
amino‐acid starvation by modulating Sec16 membrane association. EMBO J, 2011. 30(18): p.
3684‐700.
60. Ferro‐Novick, S., et al., Yeast secretory mutants that block the formation of active cell surface
enzymes. J Cell Biol, 1984. 98(1): p. 35‐43.
61. Novick, P., C. Field, and R. Schekman, Identification of 23 complementation groups required
for post‐translational events in the yeast secretory pathway. Cell, 1980. 21(1): p. 205‐15.
62. Schekman, R., et al., Yeast secretory mutants: isolation and characterization. Methods
Enzymol, 1983. 96: p. 802‐15.
63. Shaywitz, D.A., et al., COPII subunit interactions in the assembly of the vesicle coat. J Biol
Chem, 1997. 272(41): p. 25413‐6.
64. Espenshade, P., et al., Yeast SEC16 gene encodes a multidomain vesicle coat protein that
interacts with Sec23p. J Cell Biol, 1995. 131(2): p. 311‐24.
65. Gimeno, R.E., P. Espenshade, and C.A. Kaiser, COPII coat subunit interactions: Sec24p and
Sec23p bind to adjacent regions of Sec16p. Mol Biol Cell, 1996. 7(11): p. 1815‐23.
66. Long, K.R., et al., Sar1 assembly regulates membrane constriction and ER export. J Cell Biol,
2010. 190(1): p. 115‐28.
67. Bi, X., R.A. Corpina, and J. Goldberg, Structure of the Sec23/24‐Sar1 pre‐budding complex of
the COPII vesicle coat. Nature, 2002. 419(6904): p. 271‐7.
68. Kung, L.F., et al., Sec24p and Sec16p cooperate to regulate the GTP cycle of the COPII coat.
The EMBO journal, 2011. 31(4): p. 1014‐27.

181
69. Nakano, A. and M. Muramatsu, A novel GTP‐binding protein, Sar1p, is involved in transport
from the endoplasmic reticulum to the Golgi apparatus. The Journal of cell biology, 1989.
109(6 Pt 1): p. 2677‐91.
70. d'Enfert, C., et al., Sec12p‐dependent membrane binding of the small GTP‐binding protein
Sar1p promotes formation of transport vesicles from the ER. J Cell Biol, 1991. 114(4): p. 663‐
70.
71. Barlowe, C. and R. Schekman, SEC12 encodes a guanine‐nucleotide‐exchange factor essential
for transport vesicle budding from the ER. Nature, 1993. 365(6444): p. 347‐9.
72. Okamoto, M., et al., High‐curvature domains of the ER are important for the organization of
ER exit sites in Saccharomyces cerevisiae. J Cell Sci, 2012. 125(Pt 14): p. 3412‐20.
73. Bannykh, S.I., T. Rowe, and W.E. Balch, The organization of endoplasmic reticulum export
complexes. J Cell Biol, 1996. 135(1): p. 19‐35.
74. Mizoguchi, T., et al., Determination of functional regions of p125, a novel mammalian
Sec23p‐interacting protein. Biochem Biophys Res Commun, 2000. 279(1): p. 144‐9.
75. Budnik, A., K.J. Heesom, and D.J. Stephens, Characterization of human Sec16B: indications of
specialized, non‐redundant functions. Scientific reports, 2011. 1: p. 77.
76. Arimitsu, N., et al., p125/Sec23‐interacting protein (Sec23ip) is required for spermiogenesis.
FEBS Lett, 2011. 585(14): p. 2171‐6.
77. Tani, K., et al., p125 is a novel mammalian Sec23p‐interacting protein with structural
similarity to phospholipid‐modifying proteins. J Biol Chem, 1999. 274(29): p. 20505‐12.
78. Morikawa, R.K., et al., Intracellular phospholipase A1gamma (iPLA1gamma) is a novel factor
involved in coat protein complex I‐ and Rab6‐independent retrograde transport between the
endoplasmic reticulum and the Golgi complex. The Journal of biological chemistry, 2009.
284(39): p. 26620‐30.
79. Lang, M.R., et al., Secretory COPII coat component Sec23a is essential for craniofacial
chondrocyte maturation. Nat Genet, 2006. 38(10): p. 1198‐203.
80. Townley, A.K., et al., Efficient coupling of Sec23‐Sec24 to Sec13‐Sec31 drives COPII‐
dependent collagen secretion and is essential for normal craniofacial development. J Cell Sci,
2008. 121(Pt 18): p. 3025‐34.
81. Abou‐Haila, A. and D.R. Tulsiani, Mammalian sperm acrosome: formation, contents, and
function. Archives of biochemistry and biophysics, 2000. 379(2): p. 173‐82.
82. Saito‐Nakano, Y. and A. Nakano, Sed4p functions as a positive regulator of Sar1p probably
through inhibition of the GTPase activation by Sec23p. Genes to cells : devoted to molecular
& cellular mechanisms, 2000. 5(12): p. 1039‐48.
83. Rismanchi, N., R. Puertollano, and C. Blackstone, STAM adaptor proteins interact with COPII
complexes and function in ER‐to‐Golgi trafficking. Traffic, 2009. 10(2): p. 201‐17.
84. Wilson, D.G., et al., Global defects in collagen secretion in a Mia3/TANGO1 knockout mouse.
The Journal of cell biology, 2011. 193(5): p. 935‐51.

THE FACULTY OF SCIENCE,
UNIVERSITY OF COPENHAGEN
1.General information
Co-authorship statement
1/3
PhD School of Science
All papers/manuscripts with multiple authors enclosed as annexes to a PhD thesis synopsis
should contain a co‐author statement, stating the PhD student’s contribution to the paper.
PhD student Name
David Klinkenberg
Civ.reg.no. (If not applicable, then birth date)
140575-3495
E-mail
davdi.klinkenberg@bio.ku.dk
Department
Biology
Principal supervisor
2.Title of PhD thesis
Name
Lars Ellgaard
Position
Lektor
E-mail
lellgaard@bio.ku.dk
Accessory Proteins at ERES – Assembly of ER exit sites is regulated by interactions of p125A with lipid signals.
3.This co-authorship declaration applies to the following paper
Assembly of ER exit sites is regulated by interactions of p125A with lipid signals.
The extent of the PhD student’s contribution to the article is assessed on the following scale
A. has contributed to the work (0-33%)
B. has made a substantial contribution (34-66%)
C. did the majority of the work independently (67-100%).
Revised 3 January 2013

THE FACULTY OF SCIENCE,
UNIVERSITY OF COPENHAGEN
2/3
PhD School of Science
4. Declaration on the individual elements Extent (A, B, C)
1. Formulation in the concept phase of the basic scientific problem on the basis
of theoretical questions which require clarification, including a summary of
the general questions which it is assumed will be answerable via analyses or
concrete experiments/investigations.
2. Planning of experiments/analyses and formulation of investigative
methodology in such a way that the questions asked under (1) can reasonably
be expected to be answered, including choice of method and independent
methodological development.
3. Involvement in the analysis or the concrete experiments/investigation.
4. Presentation, interpretation and discussion of the results obtained in article
form.
5. Material in the paper from another degree / thesis :
Articles/work published in connection with another degree/thesis must not form part of the PhD thesis.
Data collected and preliminary work carried out as part of another degree/thesis may be part of the PhD thesis if
further research, analysis and writing are carried out as part of the PhD study.
Does the paper contain data material, which has also formed part of a previous degree / thesis
(e.g. your masters degree)
Please indicate which degree / thesis:
Percentage of the paper that is from the PhD degree work
Percentage of the paper that is from the other degree / thesis
Please indicate which specific part(-s) of the paper that has been produced as part of the PhD study:
6.Signatures of co-authors:
Date
Name Signature
C
C
C
C
Yes:
No:
%
%
X
Revised 3 January 2013