A Molecular Analysis of Opsin Integration at the

1

A Molecular Analysis of Opsin

Integration at the

Endoplasmic Reticulum

A thesis submitted to The University of Manchester for the degree of

Doctor of Philosophy in the Faculty of Life Sciences.

2005

Nurzian Ismail

Faculty of Life Sciences

The University of Manchester

2

Contents

List of Figures 6

List of Tables 10

Abbreviations 11

Abstract 12

Declaration 13

Acknowledgements 14

List of Publications 15

Chapter 1 - Introduction 16

1.1 The endoplasmic reticulum 17

1.2 ER-targeting signals 19

1.3 Targeting of polypeptide chains to the ER 23

1.4 The ER translocon 26

1.5 Translocon-associated components 29

1.5.1 The Sec62/63 complex 29

1.5.2 Signal peptidase complex 30

1.5.3 Oligosaccharyltransferase (OST) complex 31

1.5.4 TRAP 34

1.5.5 Glycoprotein specific ER chaperones 34

1.6 Structure and composition of the ER translocation site 37

1.7 The biosynthesis of polytopic membrane proteins at the ER 41

1.7.1 Targeting and insertion of polytopic membrane proteins at the

translocon 41

1.7.2 Lateral exit of TM domains into the ER membrane 44

1.7.3 Models of polytopic membrane protein integration into the ER

membrane 45

1.8 Membrane chaperones and polytopic proteins 46

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1.9 This study 47

1.9.1 Opsin as a model polytopic membrane protein 47

1.9.2 Use of site-specific cross-linking in the analysis of opsin integration 50

1.9.3 Overview 52

Chapter 2 - Materials and Methods 53

2.1 Materials 54

2.2 Site-directed mutagenesis 55

2.3 Generation of OPTM1-3PPL[cys115] and OPN/5-7[cys-null] mutants 58

2.4 Synthesis of truncated transcription templates lacking a stop codon 60

2.5 Preparation of semi-permeabilised cells 62

2.6 In vitro transcription and translation 63

2.7 Isolation of ribosome-nascent chain complexes 64

2.8 Cross-linking and modifications of nascent chains with sulphydryl specific

reagents 64

2.9 Solubilisation of ribosome-nascent chain complexes in C12E8 64

2.10 Immunoprecipitation 65

2.11 Endoglycosidase H digestion 65

2.12 SDS PAGE and sample analysis 66

Chapter 3 - The use of site-specific cross-linking approach to examine opsin

integration 67

3.1 Introduction 68

3.2 Optimisation of the experimental system 69

3.3 Cross-linking adducts are formed with glycosylated opsin chains 71

3.4 Cross-linking adduct formation is cysteine-dependent 74

3.5 Summary 78

Chapter 4 - The integration of the N-terminal region of opsin: TM1 to TM3 79

4.1 Introduction 80

4.2 TM1 is adjacent to discrete sets of ER components during its integration 82

4.3 TM1 environment is influenced by subsequent TM domains 84

4

4.4 TM2 has exited the translocon in the OP204 integration intermediate 88

4.5 TM3 is associated with the ER translocon in the OP164 integration

intermediate 90

4.6 TM3 exits the translocon upon chain extension 92

4.7 TM3 relocation is independent of the presence of subsequent TM domains 94

4.8 Nascent opsin chains are associated with a single copy of the Sec61

complex during integration 96

4.9 Summary 98

Chapter 5 - The integration of the C-terminal region of opsin: TM4 to TM7 99

5.1 Introduction 100

5.2 TM4 exits the translocon upon chain extension 100

5.3 Opsin TM5 engages the translocon at 259 residues 104

5.4 Opsin TM5 is adjacent to a PAT-10-like molecule during its integration 106

5.5 Opsin TM1 and TM5 are adjacent to a single copy of PAT-10 110

5.7 Opsin TM7 is associated with the translocon throughout opsin biosynthesis 115

5.8 Summary 118

Chapter 6 - Probing the environment of a translocating nascent opsin chain 119


6.2 AMS and QSY can modify cysteine probes in the ER lumen 121

6.3 Opsin nascent chains are modified by AMS in the absence of membranes 124

6.4 The environment of cys124 in the OP150 and OP164 integration

intermediates is altered in the presence of the ER translocon 126

6.5 Cys124 in OP150 and OP164 integration intermediate is in a hydrophobic

environment 129

6.6 The loop region C-terminal to TM3 is in a hydrophilic environment at 164

residues 131

6.7 The OP164 nascent chain represents a true integration intermediate that is

attached to the ribosome 134

6.8 Summary 136

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Chapter 7 - Discussion 137


7.2 The integration of TM1, TM2 and TM3 is a ‘co-ordinated’ process 140

7.3 An alternative analysis of TM3 environment during opsin integration 142

7.4 Opsin TM4 exits the translocon upon nascent chain extension 145

7.5 Opsin TM5 is engaged with the ER translocon throughout opsin

biosynthesis 145

7.6 Opsin TM6 and TM7 are associated with the Sec61 complex throughout

opsin biosynthesis 147

7.7 Nascent opsin chains engage a single copy of the Sec61 complex during

opsin biosynthesis 147

7.8 The possible role of PAT-10 as a TM chaperone 148

7.9 Conclusion 149

Chapter 8 - Bibliography 151

Appendices 171

Publications 176

6

List of Figures

Chapter 1

Page

Figure 1.1 A schematic representation of the secretory pathway.

18

Figure 1.2 A schematic representation of a typical cleavable ER-targeting signal sequence.

19

Figure 1.3 Topologies resulting from cleaved and uncleaved signal sequences.

20

Figure 1.4 SRP-dependent targeting of proteins destined for the ER.

25

Figure 1.5 The topology of components of the ER translocon.

27

Figure 1.6 Schematic diagram of the subunits of signal peptidase, TRAP and the oligosaccharyltransferase complex.

33

Figure 1.7 Regulation of glycoprotein folding in the ER.

36

Figure 1.8 The crystal structure of the Sec61 heterotrimer from the (a)-(b) top view and the (c) side view.

38

Figure 1.9 A schematic diagram of the putative arrangement of Sec61 complexes within the oligomer.

40

Figure 1.10 A classical model of the translocation of polytopic membrane protein into the ER membrane.

42

Figure 1.11 Schematic representation of (a) sequential integration of polytopic membrane proteins, and (b) integration of polytopic membrane proteins upon completion of translation.

45

Figure 1.12 A diagrammatic representation of bovine opsin sequence.

49

Figure 1.13 A schematic diagram of a rod photoreceptor cell.

50

Figure 1.14 Structure and chemical reaction of the homobifunctional cross-linking reagent, bismaleimidohexane (BMH).

51

Chapter 3

Figure 3.1 Rationale for HA tagging of nascent opsin chains.

70

7

Figure 3.2 Immunoprecipitation with α-HA antisera allows selection of authentic opsin chains.

72

Figure 3.3 Cross-linking products are formed with glycosylated opsin chains.

73

Figure 3.4 Formation of BMH cross-linking adducts is cysteine dependent.

76

Figure 3.5 Radiolabelled endogenous Sec61α molecules were generated during translation.

77

Figure 4.1 A diagrammatic representation of artificial opsin integration intermediates generated for site-specific cross-linking analysis of distinct TM domains.

81

Figure 4.2 BMH-mediated cross-linking of OP[cys56] integration intermediates of increasing chain length to ER components.

83

Figure 4.3 A) A schematic representation of the OPTM1PPL[cys56] polypeptide. B) BMH cross-linking of OPTM1PPL[cys56] integration intermediates of to translocon associated components.

86

Figure 4.4 A plot of the relative efficiency of cross-linking to Sec61α versus the chain length of the integration intermediate.

87

Figure 4.5 BMH cross-linking of integration intermediates with cys89.

89

Figure 4.6 BMH cross-linking of OP164 integration intermediates with cysteine probes in three different locations within TM3.

91

Figure 4.7 BMH mediated cross-linking of integration intermediates containing TM3 specific cysteine probes to ER translocon components.

93

Figure 4.8 A) A schematic representation of OPTM1-3PPL[cys115] polypeptide chain. B) BMH cross-linking of OPTM1-3PPL[cys115] integration intermediates to translocon components.

95

Figure 4.9 BMH cross-linking with OP164 and OP204 integration intermediates containing double cysteine probes.

97

Chapter 5

Figure 5.1 BMH cross-linking of OP204 integration intermediates from two different single cysteine probes within TM4.

102

8

Figure 5.2 Cross-linking of OP[cys154] integration intermediates of increasing chain lengths to translocon components.

103

Figure 5.3 BMH cross-linking of OP259 integration intermediates using different TM5 specific probes.

105

Figure 5.4 A) A schematic diagram of the OPN/5-7 polypeptide chain. B) Predicted topology of OPN/5-7 polypeptide chain. C) Cross-linking adducts are formed with glycosylated OPN/5-7 nascent chains.

107

Figure 5.5 BMH cross-linking of cys229 integration intermediates of A) normal-length opsin polypeptide chain, and B) OPN/5-7 polypeptide chain.

109

Figure 5.6 Double probe analysis of the OP304 integration intermediate.

111

Figure 5.7 BMH dependent cross-linking from single cysteine probes in opsin TM6.

113

Figure 5.8 BMH cross-linking of cys275 integration intermediates of A) normal-length opsin polypeptide chains, and B) OPN/5-7 polypeptide chains.

114

Figure 5.9 BMH mediated cross-linking from distinct cysteine probes in opsin TM7.

116

Figure 5.10 BMH cross-linking of cys287 integration intermediates of A) normal-length opsin polypeptide chain, and B) OPN/5-7 polypeptide chain.

117

Chapter 6

Figure 6.1 Structures of (a) 4-acetamido-4′-maleimidylstilbene-2-2′-disulfonic acid (AMS) and (b) QSY 9 C5-maleimide (QSY).

120

Figure 6.2 AMS and QSY modification of OP96[cys14] integration intermediates.

123

Figure 6.3 AMS modification of various OP130 to OP164 chains.

125

Figure 6.4 AMS modification of various OP130 to OP164 integration intermediates.

128

Figure 6.5 QSY modification of various OP130 to OP164 integration intermediates.

130

Figure 6.6 AMS and QSY modification of cys140 in (a) OP164 and (b) OP174 chains.

132

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Figure 6.7 AMS and QSY modification of (a) OP164[cys140] and (b)

OP174[cys140], in the presence of semi-permeabilised mammalian cells.

133

Figure 6.8 Isolation of ribosomes and associated OP164[cys124] chains.

135

Chapter 7

Figure 7.1 A working model for the integration of opsin into the ER membrane.

139

Figure 7.2 The roles of TM1 and TM3 during opsin integration.

141

Figure 7.3 Possible models of TM3 integration into a hydrophobic environment.

143

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List of Tables

Chapter 1

Page

Table 1.1 Yeast and mammalian homologues of the OST complex and their putative functions.

31

Chapter 2

Table 2.1 Primers used for the introduction of cysteine residues into opsin.

57

Table 2.2 PCR primers used in the generation of OPTM1-3PPL[cys115] and OPN/5-7 constructs.

59

Table 2.3 Primers used for the removal of cysteine residues from the preprolactin coding sequence of OPTM1-3PPL[cys115] construct.

59

Table 2.4 Primers used to generate truncated opsin transcription templates.

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Table 2.5 Primers used to generate truncated OPTM1PPL[cys56] and OPTM1-3PPL[cys115] transcription templates.

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Abbreviations

AAPs amino acid permeases

AMS 4-acetamido-4′-maleimidylstilbene-2-2′-disulfonic acid

BiP immunoglobulin heavy chain binding protein

BMH bismaleimidohexane

DMSO dimethylsulphoxide

ER endoplasmic reticulum

ERAD ER-associated degradation

HA haemagglutinin

OST oligosaccharyltransferase

PDI protein disulphide isomerase

PPL preprolactin

SDS sodium dodecyl sulphate

SPC signal peptidase complex

SRP signal recognition particle

TCA trichloroacetic acid

TM transmembrane

QSY QSY® 9 C5-maleimide

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Abstract

A major step in the biosynthesis of many membrane proteins is their insertion into the

membrane of the endoplasmic reticulum (ER). The insertion of a multi-spanning

membrane protein is a complex process since several transmembrane (TM) domains

have to be correctly integrated in order to enable its correct assembly. At present it is

unclear how the integration of multiple TM domains is co-ordinated by the ER

translocon. The aim of this study was to analyse the molecular environment of the TM

domains of a model seven TM domain protein, opsin, so as to better understand the

mechanism by which integration occurs.

For this purpose, stable ‘integration intermediates’ of defined lengths representing

distinct stages of opsin biosynthesis were generated by in vitro translation of truncated

mRNA in the presence of semi-permeabilised cells. Cysteine-mediated, site-specific

cross-linking and immunoprecipitation were employed to examine the environment of

these integration intermediates. In addition, cysteine-specific modification reagents with

different physical properties were used to investigate the environment of opsin TM3

during its insertion at the ER membrane.

Opsin TM domains exhibit unique patterns of adduct formation with the ER translocon

components, Sec61α and Sec61β. TM1 associates with the Sec61 complex at two

distinct stages during nascent chain extension, and this behaviour is dependent on the

presence of subsequent TM domains. The re-association of TM1 with the translocon

may well facilitate the co-ordinated integration of TMs 1-3 into the lipid bilayer. Opsin

TM4 exits the Sec61 complex as soon as the subsequent TM domain is synthesised,

while TM5, TM6 and TM7 remain associated with the ER translocon throughout

protein synthesis, suggesting their concerted release upon chain termination. Evidence

is provided that opsin is integrated via a single Sec61 heterotrimer, despite the fact that

the ER translocon appears to consist of multiple copies of the Sec61 complex. On the

basis of this work, a model is presented describing the complete integration of opsin at

the ER membrane.

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Declaration

No portion of the work referred to in the thesis has been submitted in support of an

application for another degree or qualification of this or any other university or other

institute of learning.

1) Copyright in text of this thesis rests with the Author. Copies (by any process)

either in full, or of extracts, may be made only in accordance with instructions

given by the Author and lodged in the John Rylands University Library of

Manchester. Details may be obtained from the Librarian. This page must form

part of any such copies made. Further copies (by any process) of copies made in

accordance with such instructions may not be made without the permission (in

writing) of the Author.

2) The ownership of any intellectual property rights which may be described in this

thesis is vested in The University of Manchester, subject to any prior agreement

to the contrary, and may not be made available for use by third parties without

the written permission of the University, which will prescribe the terms and

conditions of any such agreement.

3) Further information on the conditions under which disclosures and exploitation

may take place is available from the Head of Faculty of Life Sciences.

14

Acknowledgements

I would like to thank Samuel Crawshaw for kindly providing the OPTM1PPL[cys56]

construct and Figure 4.5.

I would like to dedicate this thesis to my parents, Ismail Ahmad and Azizah Mohamed

Amin. I am grateful to my family for the opportunities they have given me throughout

my life and would like them to know that I would not have been able to make it this far

without their enduring love, support and encouragement.

I wish to thank my fantastic supervisor, Professor Steve High, for his excellent guidance

and unwavering encouragement during my postgraduate studies in his laboratory. His

tireless optimism and enthusiasm have been both personally affirming and

professionally inspiring – I could never have wished for a better supervisor!

I would like to thank all the members of the High lab for their exceptional help, advice

and support not only in work but in other aspects of my life too. I am also grateful to the

badminton gang for making Thursdays so enjoyable.

Finally I would like to express my appreciation to all my friends here – in particular

Jifang, Bee Bee, Ray, Ay Lin, Anna, Maggie, Nafisa and Jane – for listening to all my

moans and groans! I wish to thank them for making my life in the UK so rich and

memorable and I hope we can continue our happy friendships together regardless of

future locations.

This work was funded by the Wellcome Trust.

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List of Publications

Lecomte, F.J.L., Ismail, N. and High, S. (2003). Making membrane proteins at the

mammalian endoplasmic reticulum. Biochemical Society Transactions 31: 1248-1252.

(Joint first author)

Wilson, C.M., Kraft, C., Duggan, C., Ismail, N., Crawshaw, S.G. and High, S. (2005).

Ribophorin I associates with a subset of membrane proteins after their integration at the

Sec61 translocon. J. Biol. Chem. 280: 4195-206.

Chapter 1 - Introduction

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CHAPTER 1 Introduction


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1.1 The endoplasmic reticulum

One important feature of eukaryotic cells is the presence of membrane-bound

subcellular compartments that perform various specialised functions. These

compartments are unique, having characteristic environments with different sets of

proteins. Regulating the flow of components into and out of these compartments is

crucial to the maintenance of their specific environments. Since most proteins are

synthesised in the cytosol, many must be targeted to their respective compartments

during or after their synthesis.

A major compartment in the cell is the endoplasmic reticulum (ER). The ER is

organised into a network of branching tubules and flattened sacs which are all

interconnected via a single internal space known as the ER lumen. The ER may appear

‘rough’ or ‘smooth’, depending on the presence of ribosomes on the cytosolic face of its

membrane. The ribosomes present on the rough ER identify it as a major site for protein

biosynthesis, and most of the membrane proteins found in the organelles of the

secretory pathway and the plasma membrane, together with soluble proteins destined

for secretion, are synthesised at the ER (Palade, 1975) (Fig 1.1).

Given that many proteins are synthesised at the ER, it follows that the ER is an

important site for protein folding and post-translational modification. Nascent chains

may undergo a number of covalent modifications at the ER, including the cleavage of

signal sequences, the addition of a carbohydrate group (N-linked glycosylation), the

replacement of a C-terminal transmembrane domain with a glycosylphosphoinositol

(GPI) anchor and the formation of disulphide bonds (Ellgaard & Ruddock, 2005; Mayor

& Riezman, 2004; Paetzel et al., 2002; Yan & Lennarz, 2005). In the case of N-linked

glycosylation, a carbohydrate group is added to an asparagine residue in the consensus

sequence Asn-X-Ser/Thr within a nascent chain, a process that is catalysed by the

oligosaccharyltransferase (OST) complex present in the ER membrane (Yan & Lennarz,

2005). These oligosaccharides function to facilitate protein folding and protein sorting

within the secretory pathway. The ER lumen also facilitates the formation of disulphide

bonds by providing a relatively oxidising environment and the process is catalysed by

ER-resident enzymes, including protein disulphide isomerase (PDI) and related proteins

(Ellgaard & Ruddock, 2005).


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Figure 1. 1 A schematic representation of the secretory pathway. As most protein synthesis occurs in the cytosol, targeting of proteins to their appropriate compartment is crucial. Secretory proteins and membrane proteins which are found in the organelles of the secretory pathway and at the plasma membrane, will normally transit through the endoplasmic reticulum first before reaching their final destination.

Other ER-resident proteins include molecular chaperones and folding factors, such as

calnexin, calreticulin, ERp57 and BiP (immunoglobulin heavy chain binding protein),

act to regulate the maturation and folding of the newly-synthesised proteins (Hebert et

al., 2005). Some of these ER chaperones also function as part of a ‘quality control’

machinery that disposes of misfolded or unassembled proteins. Misfolded or

unassembled proteins are retrotranslocated across the ER membrane back to the cytosol

where they are deglycosylated and polyubiquitinated before degradation (Ahner &

Brodsky, 2004). Once the proteins have attained their correct tertiary or quartenary

structures, they may be directed to other organelles in the secretory pathway, including

the Golgi apparatus, lysosomes, endosomes and secretory vesicles, or to the plasma

membrane via vesicular transport (Fig 1.1).

It is evident that before proteins can be sorted to other organelles in the secretory

pathway, they must first be targeted to the ER. Hence, the presence of targeting signals

in these proteins play a crucial role in ensuring that they are properly directed to the ER.

At the ER membrane, a specialised translocation machinery exists to facilitate the


19

passage of these proteins into and across the lipid bilayer (Alder & Johnson, 2004;

Johnson & van Waes, 1999; Lecomte et al., 2003).

1.2 ER-targeting signals

Newly-synthesised proteins are sorted to their appropriate compartment in the cell by

having distinctive signals within their sequence. Generally, an ER-targeting signal

comprises of a stretch of hydrophobic amino acids next to a short region of basic

residues (Nothwehr & Gordon, 1990). For example, secretory proteins destined for the

ER have a short, positively-charged N-terminal region (n-region), a hydrophobic core

(h-region) and a polar C-terminal region (c-region) which contains the recognition site

for signal peptidase, within their signal sequence (Fig 1.2) (Emanuelsson & von Heijne,

2001; von Heijne, 1998). The presence of such an ER-targeting signal in a protein

allows components in the cytosol or the ER membrane, such as the signal recognition

particle (SRP) and the ER translocon, to recognise it as being destined for the ER (Belin

et al., 1996; Powers & Walter, 1996).

Figure 1. 2 A schematic representation of a typical cleavable ER-targeting signal sequence. The signal sequence contains a positively-charged N-terminal region (n), a hydrophobic region (h) and a C-terminal region which contains a signal peptidase complex (SPC) cleavage site (indicated with an arrow).

Even though the ER-targeting signals of different precursors share very little sequence

identity, the most consistent feature is the presence of the central hydrophobic core (von

Heijne, 1985). The diverse signal sequences do not seem to affect the targeting

efficiency of the proteins to the ER, but may result in different rates of initiation of

nascent chain translocation (Kim et al., 2002). Despite such sequence variation, ER-

targeting signals can be swapped between substrates without any obvious consequences


20

for the function of the protein (Gierasch, 1989). Targeting usually occurs during the

translation of polypeptide chains in mammalian cells, while it may be co-translational or

post-translational in yeast cells, depending on the substrate. Once the signal sequence

has been translocated into the ER membrane, it may be cleaved or retained to function

as a transmembrane anchor for the protein (see Fig. 1.3).

Figure 1. 3 Topologies resulting from cleaved and uncleaved signal sequences. Cleavage of the signal sequence (orange) of a secretory protein would release it into the ER lumen (1) while cleavage of the signal sequence of a membrane protein would result in a type I membrane protein that is anchored via a stop-transfer sequence (brown) located after the signal sequence (2). Membrane proteins with an uncleaved signal sequence, i.e. a signal-anchor (blue), may have either a type I (3) or type II (4) orientation, depending on the properties of the hydrophobic core and the regions flanking it (see text for details). Tail-anchored proteins (5) have a type II orientation resulting from the uncleaved signal sequence (blue) near their C-termini. Polytopic membrane proteins have both signal-anchor and stop-transfer sequences, and may have either the N-terminus (6) or C-terminus (7) or both (not shown) translocated across the membrane.


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Cleavable ER-targeting signals are normally located at or towards the N-terminus of a

protein and are eventually oriented such that their N-terminus is in the cytosol when the

signal is inserted into the ER membrane. This orientation allows the cleavage of the

signal sequence by the signal peptidase complex (SPC) in the lumen to release the

signal peptide into the lipid bilayer. Secretory proteins tend to have cleavable signal

sequences as they have to be completely translocated across the ER membrane into the

lumen (Fig. 1.3). Some membrane proteins also have a cleavable signal sequence but

this is followed by another stretch of hydrophobic residues which functions as a ‘stop-

transfer’ sequence. This hydrophobic region prevents further translocation and anchors

the protein within the ER membrane. As cleavage of the signal sequence occurs on the

lumenal side of the ER membrane, membrane proteins with a cleavable signal sequence

will always have a so called type I topology with the N-terminus located in the lumen of

the ER (Fig. 1.3).

In the case of non-cleavable ER targeting signals, the signal sequence is referred to as a

signal-anchor sequence, and such signals are normally found in integral membrane

proteins (Meacock, 2000). A signal anchor sequence is responsible for both targeting

the protein to, and anchoring it in, the ER membrane. Single-spanning and multi-

spanning integral membrane proteins with a signal anchor sequence at their N-terminus

may adopt either a type I orientation, in which the N-terminus is in the lumen, or a type

II orientation, in which the N-terminus is in the cytosol (High & Laird, 1997; Meacock,

2000) (see also Fig. 1.3). Signal-anchor sequences may also be found at the C-terminus

of single-spanning integral membrane proteins. These proteins are known as tail-

anchored proteins, since the bulk of their polypeptide chain is located in the cytosol

(Fig. 1.3).

The orientation that a signal anchor sequence adopts during insertion into the ER

membrane is influenced by the distribution of positively charged side chains flanking

the central hydrophobic region (Goder & Spiess, 2001; Hartmann et al., 1989; Levy,

1996; van Geest & Lolkema, 2000). This phenomenon is sometimes referred to as the

‘positive-inside’ rule where the more positive flanking region of a signal sequence

remains in the cytosol while the less positive flanking region is translocated across the

membrane (Hartmann et al., 1989; von Heijne, 1989). Therefore, a net positive charge

towards the N-terminal region of a signal anchor sequence would result in the


22

translocation of its C-terminal flanking region and vice versa. In many cases, the

topology of a membrane protein may simply be inverted by the addition of positively-

charged residues to the appropriate flanking region (Hermansson et al., 2001; Szczesna-

Skorupa & Kemper, 1988). Whilst a cleavable signal sequence adopts a loop like

topology during translocation (c.f. Fig. 1.3), a recent study suggests that this orientation

may be achieved ‘kinetically’. Hence, the N-terminus of a signal sequence was shown

to be translocated into the ER lumen before its subsequent inversion to the opposite

orientation occurs. A more positive N-terminal region results in a faster rate of

inversion to orientate the N-terminus in the cytosol (Goder & Spiess, 2003). The basis

for the positive-inside rule for both cleavable signal sequences and signal anchors is

most likely due to interactions with the Sec61 complex of the ER translocon since

mutation of conserved residues in the Sec61 complex allowed ‘less stringent’

orientation (Goder et al., 2004).

The length and the hydrophobicity of the hydrophobic region of a signal anchor

sequence can also have a role in determining the orientation of a membrane protein, and

a longer hydrophobic segment seems to promote N-terminal, rather than a C-terminal,

translocation (Abell et al., 2002; Rosch et al., 2000). The ‘more hydrophobic’ end of the

signal sequence is also more efficiently translocated across the ER membrane (Harley et

al., 1998). Amino acid residues with greater hydrophobicity seemed to promote

translocation of the N-terminal region of the signal sequence and experiments have been

performed to rank amino acids in order of their capacity to promote N-terminal

translocation (Liu & Deber, 1998; Rosch et al., 2000). In a recent study, it was shown

that TM domain insertion may also be influenced by the position of residues within the

TM helix (Hessa et al., 2005). For example, tyrosine and tryptophan residues promote

membrane insertion when they are placed away from the centre of the helix, while a

proline residue introduced at the N-terminal end of the TM domain allow more efficient

integration.

Other factors which influence the topology of membrane proteins include, folding of the

N-terminal domain, interaction with adjacent transmembrane domains and glycosylation

(Goder & Spiess, 2001; van Geest & Lolkema, 2000). Since the N-terminal region of a

signal sequence is exposed to the cytosol first, folding of this region may affect its

ability to be translocated, thus retaining it in the cytosol. The topology of some


23

transmembrane (TM) domains is also dependent on adjacent TM domains. For example,

in the Band 3 protein, the stop-transfer sequence in the second transmembrane domain

must interact with the preceding signal anchor sequence for its integration into the

membrane. This is likely to reflect its ‘weak’ stop-transfer function, and it is capable of

being completely translocated when the loop region connecting the two TM spans is

elongated (Ota et al., 2000). Glycosylation may influence protein topology by simply

sterically trapping segments of the nascent chain in the ER lumen. In one case, when

chimeric polypeptide chains with two conflicting signal sequences were engineered with

a glycosylation site in between the signals, the nascent chains preferentially adopt a

topology in which the glycosylation site was in the ER lumen (Goder et al., 1999).

Additional trans-acting factors may also govern protein topology during translocation as

indicated by studies performed on proteins with topological heterogeneity such as the

prion protein and ductin (Dunlop et al., 1995; Hegde et al., 1998). In the case of the

prion protein, the TRAP complex (see section 1.5.3) seems to play some role in the full

translocation of the protein into the ER lumen (Fons et al., 2003).

1.3 Targeting of polypeptide chains to the ER

The ER-targeting signal of most proteins in mammalian cells is first recognised by a

ribonucleoprotein complex, consisting of one 7S RNA molecule and six polypeptides,

which is known as the signal recognition particle (SRP) (Luirink & Sinning, 2004). The

protein subunits have apparent molecular masses of 9, 14, 19, 54, 68 and 72 kDa. The

54 kDa subunit, SRP54, is the most highly conserved of the subunits (Hann et al., 1989)

and it plays a central role in the recognition of ER-targeting signals by SRP (Keenan et

al., 2001; Powers & Walter, 1996).

Three regions have been identified within the SRP54 subunit, known as the N, G and M

domains (Luirink & Sinning, 2004; Walter & Johnson, 1994). The C-terminal M

domain is rich in methionine and it interacts with both the RNA component (Römisch et

al., 1990) and the hydrophobic ER-targeting signal (High & Dobberstein, 1991). The

crystal structure of the SRP54 homologue of Thermus aquaticus, Ffh, showed that a

hydrophobic groove lined with methionine residues is present in the M domain and it

has been proposed that this could mediate the signal sequence recognition event

(Keenan et al., 1998). This region of SRP54 seems fully adapted to recognise the


24

variable h-regions that can occur within the signal sequence of different nascent chains

(Keenan et al., 1998). In addition, an arginine-rich α-helix within a helix-turn-helix

motif present in the M domain of SRP54 is implicated in binding to SRP RNA (Keenan

et al., 1998). The central G domain of SRP54 has a role in binding and hydrolysing

GTP which is essential for SRP function (Römisch et al., 1989), while the N-terminal N

domain of SRP54 subunit is thought to increase the efficiency of signal sequence

recognition (Keenan et al., 2001; Newitt & Bernstein, 1997).

Another subunit, SRP19, is essential in the assembly of SRP, and a Saccharomyces

cerevisiae homologue of SRP19, the Sec65 protein, has also been identified (Stirling &

Hewitt, 1992). SRP19 and Sec65p are involved in the recruitment of SRP54 to SRP and

hence have an important role in maintaining SRP integrity (Hann et al., 1992; Regnacq

et al., 1998; Stirling, 1999). SRP9, together with SRP14, is involved in slowing down of

the rate of nascent chain elongation by the ribosome after SRP binding to the signal

sequence. Mutations which prevent this elongation arrest decrease efficiency of nascent

chain translocation, thus implying that it is important for the function of SRP (Mason et

al., 2000). Cryo-electron microscopy structure of mammalian SRP in complex with an

active ribosome indicated that elongation arrest is achieved by direct binding of SRP to

the elongation-factor-binding site of the ribosome (Halic et al., 2004).

SRP is able to target nascent chains to the ER due to the presence of an SRP receptor in

the ER membrane. In mammalian cells, the SRP receptor consists of two components,

SRβ and SRα (Luirink & Sinning, 2004). SRα is peripherally attached to the ER

membrane on the cytosolic face by its strong interaction with SRβ which is a single-

spanning membrane protein. Like SRP54, both subunits of the SRP receptor are

GTPases (Millman & Andrews, 1997). During SRP-dependent targeting to the ER

membrane, SRP first binds to the signal sequence as soon as it emerges from the

ribosome, forming a complex that consists of SRP, the nascent chain and the ribosome.

At this stage, the SRP54 subunit of SRP is positioned very close to the exit site of the

ribosome (Halic et al., 2004; Pool et al., 2002) and translation of the polypeptide chain

is arrested. The ribosome then promotes GTP-binding by the SRP54 subunit of SRP and

this targets the complex to the ER by allowing a high affinity interaction between SRP

and its receptor in the ER membrane (Bacher et al., 1996).


25

Figure 1. 4 SRP-dependent targeting of proteins destined for the ER. SRP (in yellow) binds to the ER-targeting signal as soon as it emerges from the ribosome (stage 1). Binding of GTP by SRP subsequently allows a high affinity interaction of SRP with its receptor on the ER (stage 2, 3 and 4). This binding event releases the nascent chain from SRP and translocation through the membrane occurs (stage 5). GTP hydrolysis then allows SRP to be released from its receptor (stage 6) and it returns to the cytosol to bind to other nascent chains.

At the ER membrane, SRP54 in the complex binds to SRα, which then binds GTP.

Binding of SRP to its receptor repositions SRP54 away from the exit site of the

ribosome, possibly allowing the ribosome to bind to the ER translocon (Sec61 complex)

(Pool et al., 2002). The presence of the translocon subsequently induces the β subunit of

the SRP receptor to bind GTP, resulting in the release of the signal sequence from SRP

and initiating translocation of the nascent chain through the translocon (Fulga et al.,

2001) (Fig. 1.4). The release of the nascent chain from SRP only occurs in the presence

of a functional ER translocon, implying that an acceptor for the signal sequence is

necessary (Song et al., 2000). As SRP delivers an incomplete polypeptide chain to the

ER insertion site, the process of translocation is coupled to subsequent translation, thus

circumventing the problem of translocating large folded regions of polypeptide. GTP

hydrolysis by SRP54 and SRα subsequently allows the release of SRP from the SR

receptor thereby completing a ‘cycle’ of targeting.


26

Some proteins with classical ER-targeting signals do not seem to require SRP to be

directed to the ER, and in fact, S. cerevisiae is still viable in the absence of a functional

SRP pathway, although growth is dramatically perturbed (Walter & Johnson, 1994).

SRP-independent proteins usually possess a signal sequence with lower average

hydrophobicity (Ng et al., 1996), or other ill-defined features that confer SRP-

independent targeting (Matoba & Ogrydziak, 1998). To date, such precursors have

largely been characterised in yeast and their translocation tends to be post-translational.

This pathway involves other molecules such as the cytosolic Hsp70s and Ydj1p,

lumenal Hsp70s and an additional ER membrane protein complex, the tetrameric

Sec62/63p complex (Caplan et al., 1992; Rapoport et al., 1999). Many cytosolic

chaperones are upregulated as part of the adaptive response to the lack of an SRP

pathway consistent with their role in promoting an alternative targeting route (Mutka &

Walter, 2001).

1.4 The ER translocon

The ER translocon is composed of a heterotrimeric complex known as the Sec61

complex which is evolutionarily conserved in archea, bacteria and yeast cells (Rapoport

et al., 1996). The first component of the ER translocon, Sec61p, was initially identified

in yeast from a genetic screen for secretion mutants (Deshaies & Schekman, 1987) and

is found to be in complex with two other components, Sbh1p (Panzner et al., 1995) and

Sss1p (Esnault et al., 1993; Hartmann et al., 1994). In S. cerevisiae, two homologues of

the Sec61 complex are present, each performing slightly different functions. The

Sec61p/Sbh1p/Sss1p complex mediates co-translational translocation of nascent chains

into the ER but may additionally associate with the Sec62/63p complex to mediate post-

translational translocation of protein substrates (Panzner et al., 1995). (see also section

1.5.1). The second yeast homologue of the Sec61 complex consists of Ssh1p, a non-

essential Sec61p homologue, Sbh2p, a second Sbh1p homologue, and Sss1p (Finke et

al., 1996). This complex does not associate with the Sec62/63p complex and thus, is

proposed to be exclusively involved in co-translational translocation at the ER (Finke et

al., 1996). In bacteria and archaea, the SecY and SecE proteins are homologous to

Sec61α and Sec61γ respectively. A Sec61β homologue of archaea, Secβ, also exists but

no obvious homology was observed between Sec61β and the corresponding SecG


27

component in bacteria (Hartmann et al., 1994; Matlack et al., 1998). The bacterial

SecYEG complex and archaeal SecYEβ complex are important components of the

prokaryotic protein export machinery.

Figure 1. 5 The topology of components of the ER translocon. Sec61α and TRAM are polytopic membrane proteins with ten (confirmed) and eight (estimated) transmembrane domains respectively, while Sec61β and Sec61γ are both tail-anchored proteins. The TRAM protein is also glycosylated at an asparagine residue (shown by Y).

The major component of the mammalian Sec61 complex is the Sec61α subunit which

has 50% identity to yeast Sec61p (Görlich et al., 1992b). Sec61α is a multispanning,

non-glycosylated, integral membrane protein with ten transmembrane regions

(Wilkinson et al., 1996) (Fig. 1.5). Initial studies using photocross-linking indicated that

translocating nascent chains are in close contact with Sec61α, suggesting that it is a

component of the ER translocon (High et al., 1991; Krieg et al., 1989; Thrift et al.,

1991; Wiedmann et al., 1989). Even though Sec61α is 53 kDa in molecular mass, it was

initially detected as a 35-kDa protein when analysed by SDS-PAGE in cross-linking

studies, thus it was initially referred to as P37 and imp34 (High et al., 1991; Kellaris et

al., 1991). The Sec61α protein is essential for the translocation of both membrane and

secretory proteins across the ER membrane (Oliver et al., 1995).

Sec61α may be purified as a complex with Sec61β, the mammalian homologue of

Sbh1p, and Sec61γ, the mammalian homologue of Sss1p (Görlich & Rapoport, 1993).


28

Sec61β is a 14-kDa tail-anchored integral membrane protein whose N-terminus is in the

cytosol (Hartmann et al., 1994) (Fig. 1.5). Although S. cerevisiae Sec61β homologues,

Sbh1p and Sbh2p, are not essential for cell viability (Esnault et al., 1993; Finke et al.,

1996), in vivo studies using Drosophila showed that mutations in Sec61β are lethal and

resulted in severe defects in embryonic development (Valcarcel et al., 1999). The role

of Sec61β is still unclear but in vitro studies show that it enhances the rate of protein

translocation across the ER membrane (Kalies et al., 1998). Sec61β could also be cross-

linked to the 25-kDa subunit of the signal peptidase complex (SPC25), implying it may

play a role in the recruitment of this enzyme complex to the ER translocon (Kalies et

al., 1998). Another putative role for Sec61β is that of ribosome-binding which may

occur alongside ribosomal interactions with other components of the Sec61 complex

(Levy et al., 2001). A functional interaction between Sec61β and the nascent chain may

also be possible since it can be cross-linked to a number of translocating nascent chains

(Knight & High, 1998; Laird & High, 1997; Meacock et al., 2002).

Like Sec61β, the other component of the Sec61 complex, Sec61γ, is also a tail-anchored

integral membrane protein with a molecular mass of around 10 kDa. A single Sec61γ

homologue, Sss1p, is present in yeast and it is clearly essential for cell viability and

protein translocation, although its role is still uncertain (Esnault et al., 1993; Esnault et

al., 1994). The overexpression of Sss1p could restore translocation in a SEC61 mutant

while its depletion resulted in the accumulation of secretory and membrane proteins that

were not post-translationally modified (Esnault et al., 1993). In C. elegans, deletion of

Sec61γ gave a lethal phenotype while its mutation resulted in defects in oogenesis

(Iwasaki et al., 1996). One significant observation is that mammalian Sec61γ can

functionally replace S. cerevisiae Sss1p, indicating functional conservation of the ER

translocation complex (Hartmann et al., 1994).

Functional reconstitution of translocon components identified an additional protein that

is necessary for the translocation of several proteins across the ER (Görlich et al.,

1992b). This component is known as TRAM (translocating chain-associating membrane

protein) and it is a 34 kDa integral membrane protein believed to have eight

transmembrane regions (Fig. 1.5). To date, TRAM appears to be restricted to higher

eukaryotes and no definitive S. cerevisiae equivalent has been discovered. TRAM is

required for the efficient in vitro translocation of most, but not all, secretory and


29

membrane proteins (Görlich & Rapoport, 1993; Oliver et al., 1995; Voigt et al., 1996).

Cross-linking studies indicate that TRAM interacts primarily with the N-terminal region

preceding the hydrophobic core of the signal sequence and show that this association

occurs at early stages of the membrane translocation process (Mothes et al., 1994). The

requirement for TRAM was later shown to be dependent upon the structure of the

hydrophobic core and the length of the charged, N-terminal, region of the signal peptide

(Voigt et al., 1996).

1.5 Translocon-associated components

1.5.1 The Sec62/63 complex

The Sec62/63p complex of the ER membrane of S. cerevisiae comprises of Sec62p,

Sec63p, Sec71p and Sec72p. Sec62p is a 30-kDa integral membrane protein with two

transmembrane regions while Sec63p is a 73-kDa integral membrane protein with three

transmembrane regions (Deshaies et al., 1990). Both Sec62p and Sec63p are essential

for yeast cell viability, whilst Sec71p and Sec72p are non-essential proteins (Deshaies

& Schekman, 1989; Fang & Green, 1994; Feldheim & Schekman, 1994; Rothblatt et

al., 1989). Sec71p is a 31.5-kDa glycoprotein which spans the ER membrane once

(Fang & Green, 1994). Sec72p is a 23-kDa peripheral ER protein which interacts very

strongly with Sec71p on the cytosolic side of the membrane and it is known to bind the

signal sequence of protein precursors (Feldheim & Schekman, 1994).

The Sec62/63p complex associates with the trimeric Sec61p translocon to form a

complex of seven proteins that can mediate post-translational translocation when

purified and reconstituted into proteoliposomes (Panzner et al., 1995). Sec62p was

shown to bind this Sec complex via its cytosolic N- and C-terminal domains. It was also

shown to bind the last 14 residues of Sec63p via its N-terminal domain (Wittke et al.,

2000). In addition, the region immediately after the second transmembrane domain of

Sec62p may be important for signal sequence recognition (Wittke et al., 2000). Efficient

post-translational translocation does not only require the Sec62/63p complex, both

Kar2p (a lumenal Hsp70) and ATP are also necessary (Panzner et al., 1995). Although

Sec63p was initially characterised to be involved in post-translational translocation,

other studies have shown that it also has a role in the co-translational SRP-dependent

pathway (Willer et al., 2003; Young et al., 2001). Depletion of Sec63p from yeast cells


30

prevented the translocation of both SRP-independent and SRP-dependent precursors,

indicating the dual role of Sec63p (Young et al., 2001).

Although no such post-translational pathway has been characterised for mammalian

cells, human homologues of Sec62p and Sec63p have been identified, termed as Sec62

and Sec63 respectively (Meyer et al., 2000). The mammalian proteins are ubiquitous

and are also able to associate with the Sec61 complex. Despite the abundance of the

various subunits in the ER membrane, only low concentrations of Sec61-Sec62-Sec63

complexes were found (Meyer et al., 2000). It is not known if this complex also has a

role in post-translational protein translocation, as to date, no efficient post-translational

translocation has been observed in mammals. However, mammalian Sec62p and Sec63p

have both recently been shown to be in close proximity to membrane proteins during

their insertion at the ER membrane (Abell et al., 2003). In humans, mutations in the

SEC63 gene can give rise to autosomal dominant polycystic liver disease, potentially

via a defect in the secretion of particular proteins (Davila et al., 2004).

1.5.2 Signal peptidase complex

An important post-translational modification for a nascent chain with a cleavable signal

sequence (Fig. 1.3) is the removal of its signal peptide from the polypeptide chain after

its translocation at the ER. Cleavage of the signal sequence occurs in the ER lumen and

is mediated by the signal peptidase complex (SPC) which is composed of five

membrane proteins, named after their corresponding molecular masses: 12, 18, 21,

22/23 and 25 kDa (Evans & Blobel, 1986). SPC18, SPC21 and SPC22/23 are type II

single-spanning membrane proteins with most of their C-terminal region in the lumen of

the ER (Shelness et al., 1993), while SPC12 and SPC25 have two TM domains with

both the N- and C-termini in the cytosol (Kalies & Hartmann, 1996) (Fig. 1.6).

SPC22/23 is also singly glycosylated at its C-terminus (Fig. 1.6).

Since cleavage of the signal sequence occurs in the lumen, it is not surprising that

SPC18, SPC21 and SPC22/23 appear to have a more direct role in cleavage activity

consistent with their topologies. The yeast homologue of SPC18 and SPC21, termed

Sec11p, and the homologue of SPC22/23, Spc3p, are both essential for viability and

cleavage activity in yeast cells (Bohni et al., 1988; Meyer & Hartmann, 1997). The


31

catalytic sites of SPC reside in SPC18 and SPC21, and their cleavage activity relies on a

serine, histidine and two aspartic acid residues (VanValkenburgh et al., 1999). On the

other hand, yeast homologues of SPC12 and SPC25, known as Spc1p and Spc2p

respectively, are non-essential under normal growth conditions and do not have roles in

signal sequence cleavage (Fang et al., 1996; Mullins et al., 1996). These subunits may

have roles in modulating cleavage activity since overexpression of Spc1p can suppress

sec11 temperature-sensitive mutants while the depletion of Spc2p renders yeast cells

defective in signal sequence cleavage at a high temperature (Mullins et al., 1996).

1.5.3 Oligosaccharyltransferase (OST) complex

Yeast Mammalian Putative function(s)

Ost1p Ribophorin I Substrate binding, ER retention of complex

Ost2p DAD1 ?

Swp1p Ribophorin II ER retention of complex

Wbp1p OST48 ER retention of complex

Ost4p OST4 ?

Ost3p N33, DC2 ?

Ost6p IAP, DC2 ?

Stt3p STT3A/B Catalytic site

Ost5p ?

KCP2 ?

Table 1. 1 Yeast and mammalian homologues of the OST complex and their putative functions.

Asparagine-linked (N-linked) glycosylation occurs in the lumen of the ER and is

catalysed by a multisubunit enzyme complex known as the oligosaccharyltransferase

(OST) complex which is closely associated with the ER translocon (Chavan et al.,

2005). The OST complex catalyses the en bloc transfer of a preformed oligosaccharyl

moiety (Glc3Man9GlcNAc2) from the lipid carrier dolichyl phosphate to asparagine

within selected Asn-X-Ser/Thr consensus sequences, where X can be any residue except

for proline (Bause & Hettkamp, 1979; Yan & Lennarz, 2005). In S. cerevisiae, nine

proteins have been identified that make up the OST complex (Table 1.1). Five of these


32

proteins, Ost1p, Ost2p, Stt3p, Wbp1p and Swp1p, are essential for cell viability, while

one protein, Ost4p, is necessary for growth at 37 °C but not at 25 °C, and the other

subunits, Ost3p, Ost5p and Ost6p, are only required for optimal glycosylation activity

(refer to Knauer & Lehle, 1999 and references therein).

Several mammalian homologues of the yeast OST subunits have been identified (Yan &

Lennarz, 2005) (Table 1.1). A recent study has additionally identified two as yet

uncharacterised proteins, DC2 and KCP2, which were copurified with the mammalian

OST complex (Shibatani et al., 2005). DC2 is a ~17 kDa protein which has a weak

homology to the C-terminal region of Ost3p and Ost6p, while KCP2 is a novel ~14 kDa

protein (Shibatani et al., 2005). Of these proteins, the most conserved subunit is STT3

with more than 50% identity between yeast and mammalian homologues (Knauer &

Lehle, 1999). Photo-cross-linking studies using translocating nascent chains showed

that STT3 is in close proximity to the glycosylation consensus site (Nilsson et al.,

2003). Since cross-linking to this consensus site was abolished in the presence of a

competitive peptide substrate, STT3 became the primary candidate for the enzymatic

activity of the OST complex. Two isoforms of STT3, STT3A and STT3B, have recently

been found in human cells. These STT3 isoforms are found in different OST complexes

and have distinct catalytic activity (Kelleher et al., 2003).

Not much is known about the functions of the other subunits, but some of these proteins

are likely to have a role in peptide binding or the retention of the OST complex at the

site where it functions in the ER. Photolabelling experiments suggest that Ost1p (yeast

homologue of ribophorin I) has a possible role in substrate binding since it can bind a

I125-labelled photoreactive peptide (Yan et al., 1999). A recent study also showed that

ribophorin I can be cross-linked to a subset of newly-synthesised substrates (Wilson et

al., 2005). Ribophorin I, ribophorin II and OST48 have specific ER localisation signals

and may function to retain the complex in the ER (Fu & Kreibich, 2000).


33

Figure 1. 6 Schematic diagram of the subunits of signal peptidase, TRAP and the oligosaccharyltransferase complex. N-glycosylation sites are indicated with ‘Y’ while black rectangles represent TM domains or hydrophobic regions. (The diagram is not to scale)


34

1.5.4 TRAP

Another complex that is found associated with the ER translocon is the TRAP

(translocon-associated protein) complex. TRAP consists of four different subunits,

TRAPα, TRAPβ, TRAPγ and TRAPδ (Hartmann et al., 1993) (Fig 1.6). TRAPα was

initially thought to be an ER localised ‘signal sequence receptor’ and was consequently

called SSRα (signal sequence receptor α subunit), but depletion of SSRα did not inhibit

translocation activity at the ER (Migliaccio et al., 1991) and thus, it was renamed

TRAPα (Hartmann et al., 1993). TRAPβ (previously known as SSRβ) was initially

identified in a complex with TRAPα when the detergent Nonidet P-40 was used to

solubilise canine pancreatic microsomes (Gorlich et al., 1990), but the use of digitonin

as a ‘milder’ detergent resulted in the further identification of TRAPγ and TRAPδ

(Hartmann et al., 1993).

Cross-linking studies indicate that TRAPα is adjacent to various translocating nascent

chains (Görlich et al., 1992a), but its function is not entirely clear. A recent study using

the prion protein indicated that its proper translocation into the ER requires the TRAP

complex (Fons et al., 2003). Interestingly, the dependency of substrates on TRAP seems

to be influenced by the nature of their signal sequences. Substrates with signal

sequences which display a low efficiency in initiating translocation at the translocon are

more likely to be TRAP-dependent (Fons et al., 2003). One model suggests that TRAP

may function to stabilise the nascent chain by having an indirect effect on the translocon

structure (section 1.6), thus allowing the nascent chain to have an easier access to the

lumen, while a second model suggests that TRAP stabilises nascent chains with weak

signal sequences by interacting directly with them at the translocon.

1.5.5 Glycoprotein specific ER chaperones

The majority of proteins synthesised at the ER, including opsin, are N-glycosylated.

One of the most important functions of this modification is to promote the correct

folding of polypeptides in the ER. The addition of a large polar carbohydrate group

helps to orient the local region of the polypeptide at the surface of the protein and

decreases the likelihood of protein aggregation by increasing its solubility. In the ER,

folding of glycosylated proteins is regulated by a unique chaperone system known as


35

the calnexin-calreticulin cycle. Both calnexin and calreticulin are ER resident lectins

which interact with the glycan moiety of newly-synthesised glycoproteins, in

combination with their co-chaperone, ERp57 (Helenius & Aebi, 2004). Calnexin is a

type I single-spanning membrane protein of 65 kDa, while calreticulin is a soluble

homologue of calnexin of 46 kDa. ERp57 is a ~60 kDa thiol oxidoreductase and a

member of the protein disulphide isomerase family (Freedman et al., 1994).

When a nascent chain is first N-glycosylated, the glycan group contains three terminal

glucose residues (Fig. 1.7). Two glucose residues are successively removed from the

glycan moiety by ER glucosidases I and II to give a monoglucosylated form which is

recognised by calnexin or calreticulin (Hammond et al., 1994; Spiro et al., 1996). This

interaction is likely to be co-translational especially when the N-glycosylation site is

located towards the N-terminus of the polypeptide chain (Molinari & Helenius, 2000).

The further removal of the third glucose residue by glucosidase II prevents the

interaction of calnexin/calreticulin with the glycoprotein. If the polypeptide chain fails

to attain a native structure, it is bound by another ER lumenal enzyme, uridine

diphosphate (UDP)-glucose:glycoprotein glucosyltransferase (UGGT), which catalyses

the addition of a glucose residue back to the glycan thereby allowing

calnexin/calreticulin binding to take place again (Parodi, 2000). By recognising the

folding status of polypeptide chains, UGGT acts as a ‘folding sensor’ that forces

misfolded proteins to remain in the calnexin-calreticulin cycle.

Proteins may escape the calnexin/calreticulin cycle after the removal of a mannose

residue by another ER enzyme, α-(1,2)-mannosidase I, but its action is slow, allowing

time for the glycoproteins to undergo several rounds of the calnexin/calreticulin

mediated folding cycle (Wang & Hebert, 2003). At this stage, if the protein is properly

folded, it will be transported to the Golgi apparatus, but if it is incorrectly folded, it will

be reglucosylated by UGGT and recognised by calnexin. The misfolded protein is then

transferred to another lectin, known as EDEM (ER degradation-enhancing α-

mannosidase-like protein) which targets it to the ER-associated degradation pathway

(ERAD), where retrotranslocation of the protein back to the cytosol and degradation by

the proteasome occur (Molinari et al., 2003; Oda et al., 2003).


36

Figure 1. 7 Regulation of glycoprotein folding in the ER. Glucosidase I and II cleave two glucose residues from the oligosaccharide group, allowing calnexin or calreticulin to bind the monoglucosylated glycan group on the nascent chain. The last glucose residue is subsequently cleaved off by glucosidase II, followed by the removal of a mannose residue by α-(1,2)-mannosidase I. Properly folded glycoproteins are allowed to proceed to the Golgi, while a glucose residue is re-added onto the glycan group of polypeptide chains with non-native conformations by the action of UDP-glucose:glycoprotein glucosyltransferase (UGGT). The re-addition of a glucose residue after the action of mannosidase I results in calnexin/calreticulin binding of the nascent chain and subsequent transfer of the nascent chain to EDEM. The nascent chain is then degraded via ER-associated degradation (ERAD).


37

1.6 Structure and composition of the ER translocation site

Early structures of the ER translocon were obtained by cryo-electron microscopy and

image reconstruction techniques using purified, detergent solubilised mammalian and

yeast Sec61 complexes. These studies indicated that the Sec61 complex exists as a

cylindrical oligomeric structure consisting of three to four Sec61 heterotrimers (i.e.

Sec61α, Sec61β and Sec61γ) (Hanein et al., 1996). This ring structure has an overall

diameter of ~85 Å, a height of ~50-60 Å and an internal diameter of ~20 Å. The number

of the oligomeric particles increases when components such as the ribosome or the

Sec62/63 complex, are present, suggesting that these factors may induce the

oligomerisation of the Sec61 complexes (Hanein et al., 1996). This type of regulated

assembly is also consistent with the observation that Sec61β can only be cross-linked to

Sec61α in the presence of ribosomes (Kalies et al., 1998). The shape of the ring

structure may also be influenced by the presence of TRAP. Hence, reconstitution of

proteoliposomes containing Sec61 complexes after TRAP depletion resulted in a

structure similar to that obtained using purified Sec61 but distinct from that of Sec61

complex obtained from native membranes (Menetret et al., 2005). This implies that

TRAP may contribute to the ER translocon.

It is well-known that the ribosome has an intrinsic affinity for the translocon and it can

bind directly to the Sec61 complex (Lauring et al., 1995; Prinz et al., 2000b). In fact,

the Sec61 complex displays the properties of a major ribosomal receptor and is

protected from protease digestion in the presence of ribosomes (Kalies et al., 1994).

Limited protease digestion of the Sec61α subunit also suggests that it plays a key role in

ribosome binding (Raden et al., 2000) and such a role is entirely consistent with EM-

derived structural data (Beckmann et al., 2001). The interaction between the ribosome

and the Sec61 complex is mediated by the 28S rRNA of the large ribosomal subunit in

eukaryotes (Prinz et al., 2000a). The interaction between rRNA and the Sec61 complex

seems to be conserved since eukaryotic ribosomes can bind the prokaryotic SecYEG

complex and vice versa (Prinz et al., 2000a). The C-terminal tail or the cytoplasmic

loop 8 of Sec61α seems to be important for ribosomal interaction as proteolytic

digestion of these regions abolishes binding activity (Raden et al., 2000).


38

It was initially thought that a central cavity in the oligomeric ring structure that was

apparent in the structure deduced by cryo-EM formed the pore of the ER translocon (c.f.

Fig. 1.9). However, a recent high resolution crystal structure of the archaeal homologue

of the Sec61 complex implies a different scenario. This 3.2 Å resolution structure

suggests that one copy of the Sec61 heterotrimer is sufficient to form the active

translocation pore (Van den Berg et al., 2004) (Fig 1.8). In this model, the multi-

spanning Sec61α subunit is arranged in two halves, consisting of TM1-5 and TM6-10,

forming a clamp-shaped structure linked together by the loop between TM5 and TM6

with a central hourglass-shaped pore of ~5-8 Å (Fig. 1.8). The view that the

translocation channel is formed by one Sec61 heterotrimer is supported by the evidence

that translocating nascent chains formed the strongest adducts with residues located in

the putative pore of the comparable SecYEG complex (Cannon et al., 2005). Sec61β

makes limited contact with Sec61α and is found near the TM1-5 half, while Sec61γ

contacts TM1, TM5, TM6 and TM10 of Sec61α and clamps the two halves together. In

addition, a lateral opening between TM2-3 and TM7-8 of the Sec61α subunit is

postulated to allow the lateral exit of a signal sequence or a TM domain of a

translocating nascent chain into the phospholipid bilayer of the ER membrane (Van den

Berg et al., 2004).

Figure 1. 8 The crystal structure of the Sec61 heterotrimer from the (a)-(b) top view and the (c) side view. (a) Sec61α, Sec61β and Sec61γ subunits are represented by multi-coloured, pink and magenta strands respectively. (b) TM2a of Sec61α which acts as a plug in the pore is shown in green while all the other strands of Sec61α, β and γ are represented in white. (c) The plug (TM2a) is indicated in green and its movement during gating of the pore is shown with an arrow. The sidechains of hydrophobic residues composing the pore ring are shown in yellow. (Adapted from van den Berg et al., 2004)


39

Van den Berg et al. (2004) suggest that Sec61 complexes are arranged into an

oligomeric assembly in a back to back fashion on the basis of a 2D crystal structure of

the bacterial SecYEG translocase (Breyton et al., 2002) (see Fig. 1.9). This arrangement

is supported by several cross-linking studies with bacterial SecYEG complexes. In one

study, the N- and C-termini of two molecules of SecY (bacterial Sec61α homologue) in

a functional complex could be cross-linked together (van der Sluis et al., 2002), while in

a separate study, cross-linking experiments indicated that two SecE subunits (the

presumptive bacterial Sec61γ equivalent) are in close proximity to one another

(Kaufmann et al., 1999). This model implies that the ‘central cavity’ observed in the

ring structure seen in the low resolution EM images is simply an indentation that is

filled with lipids, rather than a water-filled channel for translocation (Dobberstein &

Sinning, 2004).

Although the crystal structure indicated the size of the translocation pore is ~5-8 Å at its

narrowest point (Van den Berg et al., 2004), other studies using fluorescent probes

suggest that the size of the pore may be larger. Fluorescence quenching agents of

different sizes were employed to establish access to fluorescent probes that were

introduced into nascent chains within the pore (Hamman et al., 1997). By determining

the largest reagent that could quench the probes, the diameter of the pore was found to

be ∼40 to 60 Å. The discrepancy in pore size may simply be a reflection of the

functional states of the translocon. The inactive translocon, as represented by the crystal

structure, may have a smaller pore which is capable of expansion in response to the

presence of a translocating nascent chain. The lifetimes of the fluorescent probes

incorporated into the signal sequence of a translocating nascent chain also indicated that

the nascent chain is in an aqueous environment during its transport across the ER

membrane (Crowley et al., 1993; Crowley et al., 1994). If the translocon pore truly has

an aqueous environment, then such an aqueous channel would allow high conductivity

when examined by electrophysiological techniques. This was indeed the case and it was

found that such a conductance was observed when empty ribosomes were bound to the

ER translocon, but the conductance was lost when the ribosomes were removed

allowing the channel to close (Simon & Blobel, 1991).


40

Figure 1. 9 A schematic diagram of the putative arrangement of Sec61 complexes within the oligomer. Individual Sec61 heterotrimers associate in a back to back manner such that the lateral opening of each Sec61 complex is facing the exterior of the structure (Adapted from Dobberstein and Sinning, 2004).

Evidence for the regulation of the translocon in order to maintain the permeability

barrier of the ER membrane suggests that, rather than being an inert structure, the

translocon is highly dynamic. Gating of the channel occurs on both cytosolic and

lumenal sides of the ER membrane. After targeting, signal sequence recognition by the

translocon occurs, leading to the formation of a seal at the ribosome-membrane junction

(Belin et al., 1996; Jungnickel & Rapoport, 1995). Some evidence suggests that the seal

may not be continuous. A three-dimensional analysis of purified Sec61 complex in

detergent in the presence of non-translating ribosomes indicated that only a single

connection exists between the complex and the ribosome (Beckmann et al., 1997). More

recently however, it was reported that there were seven connections between Sec61

complex and the ribosome with a lateral opening into the cytosol (Menetret et al.,

2005). It was speculated that these openings may allow some regions of the nascent

chain to escape into the cytoplasm during translocation. It was also proposed that the

gap between the Sec61 complex and the ribosome allows the exit of a nascent chain in

the event that the ribosome is translating a cytosolic protein (Menetret et al., 2000). It

could also be that the gap is filled with the flexible regions of the translocon

components that could not be perceived in the low resolution EM structures.

As translation continues, the pore opens to the lumen only after the length of the nascent

chain reaches about 70 residues (Crowley et al., 1994; Liao et al., 1997). During the

translocation of membrane proteins, the lumenal gate closes when the transmembrane


41

segment begins to be synthesized by the ribosome. The folding of the TM segment

within the ribosome may act as a signal to allow ribosome-induced alterations in the

conformation of the translocon (Woolhead et al., 2004). Earlier studies suggest that BiP,

a lumenal Hsp70, has a role in the closure of the lumenal end of the translocon channel

in an ATP-dependent manner (Haigh & Johnson, 2002). Pore closure requires BiP to be

ADP-bound while opening of the pore occurs only after BiP binds ATP (Alder et al.,

2005). The crystal structure of the Sec61 complex also indicates that a short TM helix

denoted TM2a may act as a ‘plug’ to block the diffusion of ions across the translocon

pore (Van den Berg et al., 2004) (Fig. 1.8(b) and (c)). In addition, a ring of hydrophobic

residues is present at the narrowest point of the translocon channel and these may form

a seal around the translocating nascent chain, thus maintaining ER membrane barrier

during substrate translocation (Van den Berg et al., 2004). Despite the need for further

studies to resolve the relative contributions of BiP and the Sec61 subunits to gating, it is

clear that the alternate opening and closing of the cytosolic and lumenal gates seem to

ensure that the permeability barrier of the ER membrane is maintained during

translocation of nascent chains.

1.7 The biosynthesis of polytopic membrane proteins at the ER

1.7.1 Targeting and insertion of polytopic membrane proteins at the translocon

The fundamental principles that underlie the biosynthesis of polytopic membrane

proteins at the ER are the same as those already described for secretory proteins and

simple membrane proteins. Hence, their targeting is SRP dependent (Friedlander &

Blobel, 1985; Wessels & Spiess, 1988) and their insertion into the ER membrane

requires the Sec61 complex (Görlich & Rapoport, 1993; High et al., 1991; Meacock et

al., 2002; Oliver et al., 1995). The biosynthesis of multi-spanning membrane proteins at

the ER is less well characterised than that of the single-spanning membrane proteins.

An early model proposed that the ribosome cycles between membrane bound and

unbound states during the biosynthesis of proteins with multiple transmembrane

domains (Blobel, 1980). In this model, the ribosome is bound to the translocon of the

ER membrane when it is synthesising domains destined for the lumen, such as regions

just after an appropriate signal sequence. The ribosome then detaches from the ER

membrane when it is translating a cytosolic segment so that this segment is synthesized

directly into the cytosol (see also Hegde & Lingappa, 1996). The next TM domain


42

would then direct the ribosome back to the translocon acting as an internal signal

sequence (Fig. 1.10).

Figure 1. 10 A classical model of the translocation of polytopic membrane protein into the ER membrane. The nascent chain is first targeted to the ER membrane via an SRP-dependent pathway (1). The ribosome binds to the translocon and translocation of the nascent chain is initiated (2). During the translation of a cytosolic domain, the ribosome detaches from the translocon such that it is synthesized directly into the cytosol (3). Synthesis of the subsequent transmembrane region directs the ribosome back to the translocon where it binds again (4). The cycle continues until the polypeptide chain is fully synthesized (5 and 6).

Experimental evidence indicates that such a model cannot account for the biosynthesis

of all polytopic membrane proteins. Cross-linking studies showed that the cytosolic

region of a nascent chain can still be cross-linked to the translocon components in the

ER membrane. Furthermore, the ribosome was not released from the membrane even

after the cytosolic domain between the transmembrane segment of the nascent chain and

the ribosome was severed by protease treatment. The region of the nascent chain that

was still attached to the ribosome after cleavage could still be cross-linked to the

translocon (Mothes et al., 1997).

One implication of the model shown in Figure 1.10 is that the orientation of the

transmembrane segments of a protein is defined by the first transmembrane sequence.

As an example, the first transmembrane segment might act as the signal sequence to

allow insertion while the second transmembrane segment acts as a ‘stop-transfer’

sequence to prevent further translocation of the nascent chain. The third transmembrane


43

region again initiates insertion while the fourth region stops translocation. Thus, the

transmembrane segments function as alternating signal and stop-transfer sequences to

eventually direct the full insertion of the polytopic polypeptide chain (Wessels &

Spiess, 1988).

In the context of this model, it is unclear if the reinitiation of subsequent signal

sequences requires SRP. By definition, the presence of a signal sequence in a

transmembrane segment would imply that SRP is needed. However, in practice, it was

shown that a short cytoplasmic segment between two transmembrane regions allowed

‘SRP-independent’ re-insertion while longer cytoplasmic regions resulted in the SRP-

dependent re-insertion of subsequent transmembrane regions (Kuroiwa et al., 1996). In

fact, not all transmembrane domains from a polytopic protein can function as a signal or

stop-transfer sequence when analysed separately (Audigier et al., 1987; Moss et al.,

1998). In some cases, it has also been found that the first transmembrane domain of a

polytopic protein does not necessarily dictate the orientation of subsequent

transmembrane domains (Sato et al., 1998). Manipulating the charge difference between

regions flanking the first TM of Glut1 glucose transporter in an attempt to invert the

topology of the entire protein resulted only in a local inversion; the topology of the

downstream TM domains of Glut1 was not affected.

A more likely scenario is that, instead of ribosomal detachment during synthesis of the

cytosolic domains of membrane proteins, the ribosome-membrane junction ‘opens’ or

‘breathes’ during translocational pausing before the next transmembrane domain is

reinitiated for insertion. Such a mechanism would allow the exit of the cytoplasmic

regions of the nascent chain into the cytosol (Beckmann et al., 2001; Hegde &

Lingappa, 1996; Menetret et al., 2000). This junction may then close during the

translocation of the next transmembrane domain. The alternate opening and closing of

the cytosolic and lumenal gates of the translocon during nascent chain translocation

maintain the permeability barrier of the ER membrane (as discussed in section 1.6).


44

1.7.2 Lateral exit of TM domains into the ER membrane

Not only is the translocon dynamic in regulating the cytosolic and lumenal gates, it must

also allow the transmembrane regions of membrane proteins to exit laterally into the

lipid bilayer. Hydrophobic regions of the stop-transfer or signal-anchor sequences of

nascent membrane proteins could be cross-linked to lipids, implying that the ER

translocon must provide access to the lipid bilayer (Heinrich & Rapoport, 2003).

Alternatively, this observation could mean that the hydrophobic transmembrane

domains can simply ‘partition’ into the lipid bilayer thereby exiting the channel

completely. It has been suggested that a role for the Sec61 complex may be to overcome

any barrier posed by the charged head groups of the phospholipids during such

‘partitioning’ of the hydrophobic transmembrane domains into the lipid bilayer

(Heinrich et al., 2000).

On the other hand, photo-crosslinking experiments have revealed that in at least one

case, a transmembrane domain of a single-spanning membrane protein goes through

three different proteinaceous environments as it exits the aqueous pore and enters the

lipid phase. Thus, the integration of a transmembrane region is not necessarily a one-

step process and it may involve several protein mediated and potentially regulated steps

(Do et al., 1996). Although this observation was seen with a single-spanning membrane

protein, the integration of TM domains of a polytopic membrane protein is likely to

occur in a similar manner. In fact, the analyses of model polytopic proteins indicate that

some TM domains remain adjacent to Sec61 components and/or TRAM for a long

period during nascent chain synthesis, even though the polypeptide chain tether from

the ribosome is long enough to allow TM diffusion out of the translocon (Meacock et

al., 2002; Sauri et al., 2005). The prolonged retention of the TM domains within the

translocon environment suggests an active regulation of TM exit by the Sec61 complex.

In addition, specific interactions between the nascent chain and the translocon are

implied by the observation that TM domains of several substrates are positioned in a

non-random manner within the translocon with respect to Sec61α and TRAM

(McCormick et al., 2003).


45

1.7.3 Models of polytopic membrane protein integration into the ER membrane

The unique feature of a polytopic protein is that several segments of the polypeptide

must at some point laterally exit the translocon, enter the lipid bilayer, and assemble to

form a functional protein. The observation that a transmembrane domain could be cross-

linked to lipids during early stages of protein biogenesis indicated that it had lateral

access to the membrane before the entire polypeptide chain had been synthesized

(Heinrich & Rapoport, 2003). The resistance of incomplete nascent chains to alkali and

urea extraction suggested that such transmembrane regions were already integrated into

the lipid bilayer before translation was terminated (Mothes et al., 1997). These data

support the view that the transmembrane segments of a polytopic protein can exit the

translocon sequentially before the entire protein has been synthesized (Fig. 1.11).

Figure 1. 11 Schematic representation of (a) sequential integration of polytopic membrane proteins, and (b) integration of polytopic membrane proteins upon completion of translation. In model (a), the transmembrane domains may exit one by one into the lipid bilayer even before translation of the full polypeptide chain has been completed. On the other hand, in model (b), integration of all the transmembrane domains occurs only after the entire polypeptide has been synthesised by the ribosome.


46

Whilst this may be true for the transmembrane domains of some membrane proteins,

other transmembrane domains appear to remain in close proximity to each other within

the environment of the translocon until translation terminates. In fact, a nascent chain

with up to five transmembrane regions could be extracted from the membrane by urea

treatment in the presence of moderate salt concentrations (Borel & Simon, 1996). This

suggests that the transmembrane regions were not integrated into the lipid bilayer, but

rather that they were stabilised by salt-sensitive, electrostatic bonds within an aqueous

pore (refer to model in Fig. 1.11(b)). It is thus possible that the translocon is able to

release transmembrane segments either sequentially, or en masse, perhaps depending on

the specific properties of the nascent chain being synthesised (High & Laird, 1997) (c.f.

Fig. 1.11(a) and (b)).

1.8 Membrane chaperones and polytopic proteins

Several ER chaperones are known to interact with secretory proteins and the lumenal

regions of membrane proteins, for example BiP and calnexin (Helenius & Aebi, 2004).

These chaperones are involved in the folding and assembly of the nascent polypeptides.

However, it is less clear whether molecular chaperones also interact specifically with

the hydrophobic domains of integral membrane proteins during their biosynthesis. To

date, a generic molecular chaperone which binds the transmembrane domains of nascent

membrane proteins has not been identified. It is possible that the exit of the

transmembrane domains of a polytopic membrane protein from the translocon is

regulated by such chaperones, and an association of this type may even define whether

their exit occurs in a sequential or en masse fashion. It could be that the Sec61 complex

and/or TRAM provide a ‘chaperone’-like function, regulating the movement of

transmembrane domains during protein biosynthesis (Do et al., 1996).

Despite the lack of evidence for a generic chaperone of transmembrane domains,

several chaperones specific for certain membrane proteins or membrane protein families

are known. One example is Shr3p, an integral membrane protein that has been shown to

be necessary for the proper folding of amino acid permeases (AAPs) which have 12 TM

domains. The insertion of AAPs into the ER membrane does not require Shr3p

(Gilstring et al., 1999), but deletion of Shr3p resulted in the aggregation of AAPs in the


47

ER (Kota & Ljungdahl, 2005), implying that Shr3p behaves as a chaperone in the

folding of AAPs. The TM domains of Shr3p are likely to be important for binding with

AAPs since a point mutation in its first TM domain abolishes this interaction (Gilstring

et al., 1999).

Other chaperones are also known to be involved in the assembly or transport of

membrane proteins rather than their integration into the ER membrane. One example is

the Vma12p and Vma22p in S. cerevisiae which form a membrane-associated complex

in the ER (Graham et al., 1998). The Vma12p/Vma22p complex interacts transiently

with a 100-kDa integral membrane protein, Vph1p, and allows it to assemble into the

Vo subcomplex of the vacuolar ATPase in the ER membrane. The Vo subcomplex later

interacts with other components to form the V-ATPase complex in vacuoles. Another

integral membrane protein which functions as a chaperone is DRiP78. DRiP78 binds to

the dopamine D1 receptor at a C-terminal hydrophobic motif, FxxxFxxxF, which is

conserved among several G protein-coupled receptors (GPCRs) (Bermak et al., 2001).

DRiP78 seems to regulate the transport of the D1 receptor away from the ER and the

export of the D1 receptor is sensitive to the levels of DRiP78 expression.

In the absence of functional Vma12p/Vma22p complex, Shr3p, and DRiP78, their

polytopic membrane protein substrates are prevented from exiting the ER, thus, these

factors play important roles in regulating protein assembly. The ‘quality control’

function of the ER ensures that only proteins which are folded and assembled correctly

may leave the ER to later compartments in the secretory pathway. Incorrectly folded

and unassembled proteins will ultimately be degraded via the ERAD pathway (Ahner &

Brodsky, 2004).

1.9 This study

1.9.1 Opsin as a model polytopic membrane protein

Opsin is a member of the guanidine-nucleotide binding protein (G-protein)-coupled

receptor (GPCR) family (Khorana, 1992) which consists of a diverse range of proteins

whose main function is to transduce an intracellular signal from an external stimuli.

Opsin has the typical topology of a GPCR; that is seven transmembrane domains with

an extracellular N-terminus and an intracellular C-terminus (Fig. 1.12). In bovine opsin,


48

several of its 348 amino acids undergo covalent modification during its biosynthesis.

Two asparagine residues near the N-terminus, Asn-2 and Asn-15, are glycosylated

(Hargrave, 1977), while two cysteines near the C-terminus, Cys-322 and Cys-323, are

palmitoylated (Ovchinnikov Yu et al., 1988). A disulphide bridge is also present

between cysteines 110 and 187 (Karnik & Khorana, 1990). Additionally, the functional

form of opsin, rhodopsin, is only generated when the chromophore 11-cis-retinal (a

derivative of vitamin A) is linked to lysine 296 of opsin (Hargrave et al., 1983).

Rhodopsin is one of the best characterised GPCRs and is currently the only GPCR with

a high resolution crystal structure, solved to 2.8 Å (Palczewski et al., 2000).

Rhodopsin is found in the plasma membrane and disc membranes of rod photoreceptor

cells (Fig. 1.13) in the retina of the eye and is responsible for peripheral and dim light

vision. A large number of mutations, most being point mutations, in the rhodopsin gene

result in an autosomal dominant disease known as retinitis pigmentosa, characterised by

the patients having tunnel vision and night-blindness. Approximately 100 mutations in

the rhodopsin gene have been identified to cause this disease and most of these

mutations affect the folding or trafficking of opsin (Kennan et al., 2005). Studies using

rhodopsin mutants expressed in cultured mammalian cells have allowed these mutants

to be categorised into two broad classes based on the intracellular fates of these mutant

rhodopsin molecules (Sung et al., 1991; Sung et al., 1993). Class I mutants are

trafficked to the plasma membrane and are functional when reconstituted with 11-cis-

retinal (Sung et al., 1991), but these mutants are defective in transport to or retention in

the rod outer segment (Sung et al., 1994). These class I mutations tend to be clustered in

the first TM domain or at the C-terminus of opsin (Deretic et al., 1996; Sung et al.,

1993). Most of the rhodopsin mutations belong to class II in which the folding or

stability of the mutant proteins are affected. As a consequence, these mutants tend to

accumulate in the ER and fail to associate with 11-cis-retinal (Sung et al., 1991; Sung et

al., 1993). Unlike class I mutations, class II mutations are generally found in TM and

extracellular domains of opsin.

During the biosynthesis of opsin at the ER membrane, the first TM domain functions as

an uncleaved signal anchor sequence which directs targeting and translocation of the N-

terminus into the ER lumen. SRP is required to direct nascent opsin chains to the ER

membrane (Friedlander & Blobel, 1985), and it was shown that some of the opsin TM


49

domains can behave as independent signal sequences and stop transfer sequences

(Audigier et al., 1987; Friedlander & Blobel, 1985). In Drosophila, the biosynthesis of a

subclass of rhodopsin isoforms involves a peptidyl-prolyl cis-trans isomerase, NinaA,

which is necessary for the proper folding and/or export of rhodopsin from the ER

(Baker et al., 1994; Colley et al., 1991; Shieh et al., 1989). NinaA forms a stable

complex with rhodopsin in vivo and its C-terminal tail is likely to be involved in this

interaction (Baker et al., 1994). Mutations in the ninaA gene result in the accumulation

of opsin at the ER and prevent opsin export to the plasma membrane (Colley et al.,

1991).

Figure 1. 12 A diagrammatic representation of bovine opsin sequence. Asparagine-linked glycan groups at residues 2 and 15 are indicated with ‘Y’, while the disulphide bond between cysteines 110 and 87 is shown as a blue dotted line. Palmitoyl groups linked to cysteines 322 and 323 are shown in magenta.


50

Figure 1. 13 A schematic diagram of a rod photoreceptor cell. The regions of the rod photoreceptor cell are marked as indicated.

1.9.2 Use of site-specific cross-linking in the analysis of opsin integration

In this study, site-specific, thiol-mediated cross-linking was utilised to examine the

proteinaceous environment of specific regions of opsin. This approach has been

extensively used to examine the translocation of polypeptide chains at the ER

membrane (Abell et al., 2003; Laird & High, 1997; Meacock et al., 2002), most often

using nascent chains which are still attached to the ribosome to provide a ‘snapshot’ of

the insertion process (Martoglio & Dobberstein, 1996). Translation of the opsin

polypeptide chain was halted at defined chain lengths by the truncation of the encoding

mRNA causing the resulting nascent chain to remain attached to the ribosome as a

peptidyl-tRNA species. When such truncated mRNAs are translated in the presence of

ER membranes, the resulting ribosome-nascent chain complexes are targeted to the ER

and become trapped in the translocon as stable ‘integration intermediate’ which are

believed to represent a particular stage of opsin biosynthesis (Gilmore et al., 1991).

When a suitable cross-linking reagent is added at this stage, the integration

intermediates can be covalently attached to adjacent ER components and subsequently

analysed by immunoprecipitation and SDS-PAGE.

The cross-linking reactions used during this study rely exclusively on the use of a

homobifunctional cross-linking reagent, bismaleimidohexane (BMH), which has two

maleimide groups that can react with two free sulphydryl groups of cysteine side chains

(Fig 1.14). BMH is able to diffuse into the lipid bilayer of the ER membrane, thus


51

allowing cross-linking of nascent chains with adjacent proteins even when the reactive

groups are located within a hydrophobic environment. BMH has a flexible spacer arm

of 16 Å, and thus can only react with available sulphydryl groups that are within a

limited proximity of each other. In order to confer site-specificity to the cross-linking

reaction, it is important that the opsin integration intermediate contains only one

cysteine residue which acts as the sole target for the reaction. In this way, any cross-

linking products formed can be attributed to a particular region of the nascent chain.

Cross-linking partners of each integration intermediate were identified by

immunoprecipitation with antisera specific for components of the ER translocon. By

performing site-specific cross-linking with opsin integration intermediates of increasing

nascent chain lengths, the proteinaceous environment of opsin could be determined at

each stage of opsin biosynthesis, allowing a model of opsin integration to be

constructed.

Figure 1. 14 Structure and chemical reaction of the homobifunctional cross-linking reagent, bismaleimidohexane (BMH).


52

1.9.3 Overview

The aim of this study is to examine the molecular environment of specific regions of

opsin, in particular, its TM domains, during its biosynthesis at the ER. This information

will be used to try to understand the mechanism by which polytopic membrane proteins

are integrated into the phospholipid bilayer of the ER membrane. Opsin is a suitable

model protein because of the availability of a high resolution crystal structure for the

wild type molecule and a good monoclonal antibody specific to its N-terminus. The

molecular environment of all seven TM domains of opsin was characterised during the

course of this study, with specific focus on opsin TM3 to TM7 which had not been

studied before in any detail (Meacock et al., 2002). The influence of later TM domains

on the lateral exit of TM1 and TM3 at the ER translocon was investigated, and the

environment of TM3 during its movement from the ribosomal exit tunnel to the ER

translocon was also probed. These data were used to build a model describing the

molecular mechanisms that underlie opsin integration.

Chapter 2 – Materials & Methods

53

CHAPTER 2 Materials and Methods


54

2.1 Materials

The plasmid encoding a cysteine-null version of the opsin coding region in the

pGEM3Z vector was previously constructed by Suzanna Meacock (Meacock, 1999).

The cross-linking reagent, bismaleimidohexane (BMH), was purchased from Perbio

Science (Northumberland, UK). The sulphydryl specific modification reagents, 4-

acetamido-4’-maleimidylstilbene-2-2’-disulfonic acid (AMS) and QSY® 9 C5-

maleimide (QSY), were purchased from Molecular Probes (Leiden, The Netherlands).

Restriction ezymes and endoglycosidase H were obtained from New England BioLabs

(Hitchin, UK). The QuikChange mutagenesis kit and the competent XL1-Blue E. coli

cells were from Stratagene (Cambridge, UK). The QIAprep Spin Miniprep kit, the

QIAquick PCR purification kit and the RNeasy mini kit were obtained from QIAGEN

(Crawley, UK). The BigDye terminator cycle sequencing ready reaction was purchased

from Applied Biosystems (Warrington, UK).

T7 RNA polymerase, SP6 RNA polymerase, transcription buffers, rNTPs, RNasin

ribonuclease inhibitor, amino acids, RNase A and nuclease-treated rabbit reticulose

lysate were obtained from Promega (Herts, UK). The cap analogue m7G(5’)ppp(5’)G

was obtained from New England BioLabs (Hitchin, UK). The aurintricarboxylic acid

(ATCA), cycloheximide and phenylmethylsulfonyl fluoride (PMSF) were purchased

from Sigma (Gillingham, UK). Easytag L-[35S]-methionine was from NEN Du Pont

(Stevenage, UK). All reagents for cell culture were obtained from Invitrogen (Paisley,

UK). Digitonin was purchased from Merck Biosciences (Nottingham, UK). All other

chemicals were analytical grade or better and were obtained from BDH/Merck (Poole,

UK) and Sigma (Gillingham, UK).

The mouse monoclonal anti-haemagglutinin (HA) antibody was a gift from Dr. I.

Hagan, Paterson Laboratories, Manchester, and rabbit polyclonal antibodies specific for

the Sec61α and Sec61β subunits were kindly provided by Prof. R. Zimmermann,

University of the Saarland, Germany. The mouse monoclonal antibody specific for the

N-terminal region of opsin was purified from a hybridoma line originally supplied by

Dr. P. Hargrave, University of Florida, USA (Adamus et al., 1991).


55

2.2 Site-directed mutagenesis

The plasmid containing a cysteine-null version of the opsin-coding region in the

pGEM3Z vector has been previously described (Meacock et al., 2002). This construct

was the starting point for the introduction of single cysteine codons by site-directed

mutagenesis. Unique cysteine residues were introduced at positions 107, 115, 124, 132,

140, 154, 165, 204, 217, 229, 254, 275, 287 and 308 of the resulting opsin chains. The

mutagenesis was performed using the PCR-based QuikChange mutagenesis kit

(Stratagene). A set of complementary primers was designed to incorporate a cysteine at

a designated amino acid position for each opsin mutant (Table 2.1). For the PCR

reaction, the following solutions were mixed: 5 µl of 10x reaction mixture (100 mM

KCl, 100 mM (NH4)2SO4, 200 mM Tris-HCl pH 8.8, 20 mM MgSO4, 1 % Triton X-

100, 1 mg/ml nuclease-free bovine serum albumin), ∼ 200 ng of DNA template, 125 ng

of forward and reverse primers, 1 µl of 2.5 mM dNTP mix, 2.5 U of Pfu Turbo DNA

polymerase and water to a final volume of 50 µl. The PCR conditions were: initial

denaturation (95 °C, 30 seconds) for 1 cycle, then denaturation (95 °C, 30 seconds),

annealing (55 °C, 1 minute) and extension (68 °C, 10 minutes) for 16 cycles.

After the PCR reaction, the mixture was incubated at 37 °C with the restriction enzyme

DpnI (10 U) for 2 hours, which specifically cleaves -GA↓TC- where the adenine

residue is methylated in parental DNA strands, leaving only non-methylated DNA with

the incorporated mutation. The remaining plasmid DNA was purified by ethanol

precipitation using ∼2.5 µg of glycogen as a carrier. The resulting DNA was then

transformed into competent XL1-Blue bacterial cells (Stratagene). Up to 5 µl of DNA

solution was added to 50 µl of competent XL1-Blue cells and the samples left on ice for

30 minutes. The cells were heat-shocked for 45 seconds at 42°C in a waterbath, and

transferred back to ice. Luria-Broth (LB) (500 µl) was then added to the mixture and the

tube was incubated at 37 °C for 30 minutes. The cells (100 µl) were plated onto LB

plates supplemented with ampicillin (100 µg/ml) and incubated at 37 °C overnight.

Random colonies were picked from the LB plate of transformed XL1-Blue cells and

cultured in LB (2 ml) with ampicillin (100 µg/ml) at 37 °C overnight. The cells were

harvested by centrifugation at full speed (∼16,000g) in a microfuge for 20 seconds at


56

room temperature and the plasmid DNA was extracted from the cells using a QIAprep

Spin Miniprep kit (QIAGEN). The resulting DNA was used for a PCR-based

sequencing reaction by mixing: 4 µl of BigDye terminator cycle sequencing ready

reaction version 1.1 (Applied Biosystems), 0.75 µl of forward and reverse primers (∼3.2

µM), 2 µl of plasmid DNA and water to a final volume of 15 µl. The forward primer

used was the pG3Z–160T7 primer which was complementary to the DNA sequence 160

bases upstream of the T7 promoter, while the reverse primer used was complimentary to

the SP6 promoter. The sequence of the pG3Z-160T7 primer is 5’-

GGGCCTCTTCGCTATTACGC-3’ whilst the sequence of the SP6 primer is 5’-

TATTTAGGTGACACTATAG-3’. The PCR conditions were: initial denaturation (96

°C, 2 minutes) for 1 cycle, then denaturation (96 °C, 30 seconds), annealing (50 °C, 15

seconds) and extension (60 °C, 4 minutes) for 29 cycles. The PCR products were then

subjected to ethanol precipitation and left in pellet form for sequencing (Sequencing

Facility, Faculty of Life Sciences, The University of Manchester). All point mutations

were confirmed by DNA sequence analysis prior to further use.

Other opsin point mutants used in this study had been previously constructed, including

OP[cys14], OP[cys56] and OP[cys87] (Meacock, 1999). The opsin-coding regions of

these mutants are present in the PGEM3Z vector, except for OP[cys14] which is present

in pZEOSV2(+) vector. For the construction of double cysteine opsin mutants, plasmids

containing OP[cys56] and OP[cys87] were used as the templates for the introduction of

a cysteine residue at position 115 by site-directed mutagenesis.


57

Table 2.1 Primers used for the introduction of cysteine residues into opsin.

Mutagenic primer Sequence of primer (5’ to 3’)

OP107C.1 GGATACTTCGTCTTTGGGTGCACGGGCGGCAACCTG OP107C.2 CAGGTTGCCGCCCGTGCACCCAAAGACGAAGTATCC

OP115C.1 GGCAACCTGGAGGGCTGCTTTGCCACCCTGGGC OP115C.2 GCCCAGGGTGGCAAAGCAGCCCTCCAGGTTGCC

OP124C.1 CACCCTGGGCGGTGAAATTTGCCTGTGGTCCTTGGTGGTCCTG OP124C.2 CAGGACCACCAAGGACCACAGGCAAATTTCACCGCCCAGGGTG

OP132C.1 GTCCTTGGTGGTCCTGTGCATCGAGCGGTACGTG OP132C.2 CACGTACCGCTCGATGCACAGGACCACCAAGGAC

OP140C.1 GAGCGGTACGTGGTGGTGTGCAAGCCCATGAGCAACTTCCGC OP140C.2 GCGGAAGTTGCTCATGGGCTTGCACACCACCACGTACCGCTC

OP154C.1 GGGGAGAACCACGCCTGCATGGGCGTCGCCTTC OP154C.2 GAAGGCGACGCCCATGCAGGCGTGGTTCTCCCC

OP165C.1 CACCTGGGTCATGGCTTGCGCCGGTGCCGCGCCC OP165C.2 GGGCGCGGCACCGGCGCAAGCCATGACCCAGGTG

OP204C.1 CCAACAATGAGTCGTTCTGCATCTACATGTTCGTGG OP204C.2 CCACGAACATGTAGATGCAGAACGACTCATTGTTGG

OP217C.1 CATCATCCCCCTGTGTGTCATATTCTTCGGCTACGGG OP217C.2 CCCGTAGCCGAAGAATATGACACACAGGGGGATGATG

OP229C.1 GGGCAGCTGGTGTTCTGCGTCAAGGAGGCGGC OP229C.2 GCCGCCTCCTTGACGCAGAACACCAGCTGCCC

OP254C.1 GGAGGTCACCCGCATGTGCATCATCATGGTCATCGC OP254C.2 GCGATGACCATGATGATGCACATGCGGGTGACCTCC

OP275C.1 GGGGTGGCGTTCTACTGCTTCACCCATCAGGG OP275C.2 CCCTGATGGGTGAAGCAGTAGAACGCCACCCC

OP287C.1 CTTTGGCCCCATCTGCATGACCATCCCGGC OP287C.2 GCCGGGATGGTCATGCAGATGGGGCCAAAG

OP308C.1 CCCCGTCATCTACATCTGCATGAACAAGCAGTTCCGG OP308C.2 CCGGAACTGCTTGTTCATGCAGATGTAGATGACGGGG

The suffix ’.1’ indicates a forward primer, while ‘.2’ indicates a reverse primer.


58

2.3 Generation of OPTM1-3PPL[cys115] and OPN/5-7[cys-null] mutants

Regions coding for opsin and preprolactin were fused to form the OPTM1-3PPL

chimera with a cysteine residue at position 115. Double-stranded DNA fragments

coding for a region of opsin with cys115 (residues 1 to 142, OPTM1-3) were generated

by PCR using the forward primer OP56C(1-70)-FOR and the reverse primer

PPL/OPTM3 (see Table 2.2 for the sequences of the primers). Similarly, double-

stranded DNA fragments coding for a region of preprolactin (residues 31 to 229, PPL)

were generated by PCR using the forward primer OPTM3/PPL and the reverse primer

PPL(31-229)-REV (Table 2.2). The PCR conditions were: initial denaturation (95 °C, 1

minutes) for 1 cycle, then denaturation (95 °C, 45 seconds), annealing (50 °C, 45

seconds) and extension (72 °C, 2 minutes) for 30 cycles. A second round of PCR using

annealed fragments of OPTM1-3 and PPL as templates with OP56C(1-70)-FOR primer

and PPL(31-229)-REV primer, gave DNA products of the OPTM1-3PPL chimera that

were subsequently cloned into the pSPUTK vector. Site-directed mutagenesis (section

2.2) was employed to replace the first three cysteine residues of the PPL coding region

within OPTM1-3PPL[cys115] with glycine residues. Complementary primers used for

the removal of these cysteine residues are shown in Table 2.3. OPTM1PPL[cys56] was

kindly provided by Samuel Crawshaw.

An opsin mutant with residues 36 to 194 deleted was generated to form the OPN/5-7

construct using a one-step PCR approach. The reverse primer OP35-REV defines the 5’

end of the region to be deleted while the forward primer defines the 3’ end of the region.

The sequences of the primers are shown in Table 2.2. A PCR reaction using these

primers resulted in the amplification of the entire plasmid which lacks the coding region

of residues 36 to 194 of opsin. The restriction enzyme DpnI was added to digest the

methylated parental DNA strand. The linear PCR products were ligated using Rapid

DNA Ligation kit (Stratagene) according to the manufacturer’s manual and transformed

into competent XL1-Blue competent cells. Extracted DNA was analysed by sequencing

as before (section 2.2). Single cysteine residues were introduced into the opsin TM

domains by site-directed mutagenesis using complementary primers as shown in Table

2.1.


59

Table 2.2 PCR primers used in the generation of OPTM1-3PPL[cys115] and OPN/5-7 constructs.

PCR primer Sequence of primer (5’ to 3’)

OPTM3/PPL CGGTACGTGGTGGTGGGCAAGCCCACCCCCGTCTGTCCCAATGGGCC

PPL/OPTM3 GGCCCATTGGGACAGACGGGGGTGGGCTTGCCCACCACCACGTACCG

OP56C(1-70)-FOR GCGAGATCTACCATGAACGGGACCGAGGGC

PPL(31-229)-REV GCGAGATCTTTAGCAGTTGTTGTTGTAGATG

OP35-REV *P-CCATGGCTCCGCCAGGTAGTACTGCGGGGC

OP195-FOR *P-CACGAGGAGACCAACAATGAGTCGTTCGTC

* ‘P’ indicates that the primers are phosphorylated.

Table 2.3 Primers used for the removal of cysteine residues from the preprolactin coding sequence of OPTM1-3PPL[cys115] construct.

Mutagenic primer Sequence of primer (5’ to 3’)

OP3PPLC34G.1 CCCACCCCCGTCGGTCCCAATGGG

OP3PPLC34G.2 CCCATTGGGACCGACGGGGGTGGG

PPLC41G.1 GGGCCTGGCAACGGCCAGGTATCC

PPLC41G.2 GGATACCTGGCCGTTGCCAGGCCC

PPLC88G.1 GCCCTCAACAGCGGCCATACCTCCTCC

PPLC88G.2 GGAGGAGGTATGGCCGCTGTTGAGGGC

The suffix ’.1’ indicates a forward primer, while ‘.2’ indicates a reverse primer.


60

2.4 Synthesis of truncated transcription templates lacking a stop codon

PCR was used to generate truncated sections of both the opsin-coding region and its

derivatives so as to provide DNA templates for in vitro transcription (see Laird and

High, 1997). The following solutions were mixed: 10 µl of 2.5 mM dNTPs, 10 µl of 10x

PWO reaction buffer (100 mM Tris-HCl pH 8.85, 250 mM KCl, 50 mM (NH4)2SO4, 20

mM MgSO4), 100 pmol each of forward and reverse primers, 5 U PWO DNA

polymerase and ∼1 µg of plasmid DNA and water to a final volume of 100 µl. For

opsin-coding regions present in PGEM3Z vector, the forward primer used was always

the pG3Z–160T7 primer (refer to section 2.2 for sequence) while the reverse primer

used depended on the length of truncated mRNA required (Table 2.4). For the synthesis

of truncated OPTM1PPL[cys56] and OPTM1-3PPL[cys115] transcription templates

from the pSPUTK vector, the forward primer used was SK-SP6, while the reverse

primer used depended on the length desired (shown in Table 2.5). The synthesis of

OP96[cys14] (in vector pZEOSV2(+)) uses the forward primer pZEO-160T7, which is

complementary to the sequence 160 bases upstream the T7 promoter in the

pZEOSV2(+) vector, and the reverse primer OP87HA-REV (Table 2.4). The sequence

of the SK-SP6 primer is 5’-CCAGAAACTCAGAAGGTTCG-3’ while the sequence of

the pZEO-160T7 primer is 5’-CCAGTTCCGCCCATTCTCCG-3’.

The PCR conditions used were: initial denaturation (94 °C, 4 minutes) for 1 cycle, then

denaturation (94 °C, 1 minute), annealing (60 °C, 1 minute) and extension (72 °C, 1

minute) for 35 cycles, and the final extension (72 °C, 10 min) for 1 cycle. The DNA

obtained was purified using QIAquick PCR purification kit (QIAGEN), and then treated

with 20 U of DpnI to remove the methylated parental DNA template. The QIAquick

PCR purification kit was again used to purify the DNA after the DpnI treatment.

Table 2.4 Primers used to generate truncated opsin transcription templates. Nascent

chain length

Truncation primer Sequence of primer (5’ to 3’)

96 OP87HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTAGACCATGAAGAGGTCGGCCAC

130 OP121HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTAACCGCCCAGGGTGGCAAAGAAGCC


61

140 OP131HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTACAGGACCACCAAGGACCACAG

150 OP141HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTACTTGCCCACCACCACGTACC

164 OP155HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTACATGATGGCGTGGTTCTCCC

174 OP165HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTACAGAGCCATGACCCAGGTG

204 OP195HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTAGTGGGGCGTGTAGTAGTCAATC

259 OP250HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTAGACCTCCTTCTCGGCCTTCTG

304 OP295HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTAGGCAAAGAAAGCCGGGATGG

339 OP330HA-REV

AGCGTAGTCTGGGACGTCGTATGGGTAGTCACCCAGCGGGTTCTTGC

357 OP348HA-REV AGCGTAGTCTGGGACGTCGTATGGGTAGGCAGGCGCCACCTGGC

Table 2.5 Primers used to generate truncated OPTM1PPL[cys56] and OPTM1-3PPL[cys115] transcription templates. Nascent

chain length*

Truncation primer Sequence of primer (5’ to 3’)

TM1 109 OPPPL109HA-REV AGCGTAGTCTGGGACGTCGTATGGGTAATGGATGTAGT

GGGACACCATGACTGCCCG

TM1 130 OP1PPL121HA-REV AGCGTAGTCTGGGACGTCGTATGGGTAGGTAATGAACC

CTTTGCCCTGGG

TM1 150 OPPPL150HA-REV AGCGTAGTCTGGGACGTCGTATGGGTATTGTTCTTTATC

TTCCGGGGTAGG

TM1 164 OP1PPL155HA-REV AGCGTAGTCTGGGACGTCGTATGGGTAAAGAATCAAGC

TCATAAGGACTTC

TM1 204 OP1PPL195HA-REV AGCGTAGTCTGGGACGTCGTATGGGTAAAGTCGTTTGTT

TTCTTCCTCAATCTC

TM1 259 OP1PPL250HA-REV AGCGTAGTCTGGGACGTCGTATGGGTACTTGCTTGAATC

CCTGCGCAGGCC

TM3 164 OP3/PPL155HA-REV AGCGTAGTCTGGGACGTCGTATGGGTATACCTGGCCGTT

GCCAGGCCCATTGGG

TM3 204 OP3/PPL195HA-REV AGCGTAGTCTGGGACGTCGTATGGGTACATGGTAATGA

ACCCTTTGCC

* The prefix ‘TM1’ indicates truncation primers for OPTM1PPL[cys56] while ‘TM3’ indicates truncation primers for OPTM1-3PPL[cys115].


62

2.5 Preparation of semi-permeabilised cells

The human HT-1080 fibrosarcoma cell line (European Collection of Cell Cultures,

Salisbury, UK) was cultured with minimal essential medium (MEM) with Earle’s salts,

supplemented with 1/100 volume 200 mM L-glutamine, 1/100 volume 100 mM sodium

pyruvate, 1/100 volume MEM non-essential amino acids, 1/10 volume foetal calf serum

and 7/500 volume MEM vitamins solution. The cells were grown to ∼90 % confluence

in 75 cm2 flasks before they were used. The cells in the flasks were washed twice with

phosphate-buffered saline (PBS) and detached by incubation with 3 ml of Trypsin-

EDTA (0.05 % w/v Trypsin, 0.53 mM EDTA.4Na). The action of trypsin was inhibited

by the addition of 4 ml of KHM buffer (110 mM potassium acetate, 2 mM magnesium

acetate, 20 mM HEPES pH 7.2) with Soybean Trypsin Inhibitor at 100 µg/ml.

The cells were semi-permeabilised as previously described by Wilson et al (1995) using

the same procedure. Cells were pelleted in 15 ml polypropylene tubes at 240g for 3

minutes at 4 °C. The cell pellet was resuspended in 4 ml of KHM buffer containing

digitonin (40 µg/ml). Samples were left on ice for 5 minutes to allow permeabilisation

of the cells, 10 ml of KHM buffer was added to the tube to dilute the digitonin, and the

cells pelleted again at 240g for 3 minutes at 4 °C. The pellet was resuspended in 5 ml of

HEPES buffer (50 mM potassium acetate, 90 mM HEPES pH 7.2) and the cells left on

ice for 5 minutes to recover. The tube was spun as before and the permeabilised cells

were resuspended in 500 µl of KHM buffer. The suspension was transferred to a

microcentrifuge tube and spun at ∼16,000g for 10 seconds before the pellet was

resuspended in 100 µl of KHM buffer. 1 µl of 0.1 M calcium chloride was added and

0.1 U of calcium-dependent micrococcal nuclease was added to the cell suspension and

the sample incubated at room temperature for 12 minutes. The action of micrococcal

nuclease was then inhibited by the addition of 1 µl of 0.4 M EGTA. The sample was

spun again at ∼16,000g for 10 seconds and the cell pellet resuspended in KHM buffer to

give a concentration of ∼0.5 x 105 cells/µl.


63

2.6 In vitro transcription and translation

For in vitro transcription of opsin constructs in PGEM3Z vector, the following solutions

were mixed in a tube: 20 µl of 5x TSC buffer (200 mM Tris-HCl pH 7.9, 30 mM

MgCl2, 10 mM spermidine, 50 mM NaCl), 10 µl of 100 mM DTT, 4 µl of 25 mM

rNTPs, 5 µl of 10 mM m7G(5’)ppp(5’)G cap analogue, 55 µl of DNA template (from

PCR), 80U of RNAsin and 80U of T7 RNA polymerase. In vitro transcription of opsin

coding regions in pSPUTK vector uses 80U of SP6 RNA polymerase with the addition

of the following: 10 µl of 10x SP6 RNA polymerase buffer (400 mM Tris-HCl pH 7.9,

60 mM MgCl2, 100 mM dithiothreitol, 20 mM spermidine), 4 µl of 25 mM rNTPs, 5 µl

of 10 mM m7G(5’)ppp(5’)G cap analogue, 55 µl of DNA template (from PCR) and 80U

of RNAsin and H2O to a final volume of 100 µl. The sample was incubated at 37 °C for

2 hours and the RNA obtained was purified using an RNeasy mini kit (QIAGEN) and

stored at -80°C.

All in vitro translations were performed in the presence of semi-permeabilised cells,

unless otherwise specified. An initial mixture of 14 µl nuclease-treated rabbit reticulose

lysate, 0.5 µl 19 amino acid mix without methionine (each at 1mM), 1.5 µl 35S-

methionine (11µCi/µl) and 4 µl semi-intact cells, were incubated at 30 °C for 3 minutes.

2 µl of RNA transcript was added and the mixture incubated at 30 °C for 15 minutes.

Further initiation of translation was inhibited by the addition of aurintricarboxylic acid

(ATCA) to 0.1 mM with an incubation of 10 minutes at 30 °C.

For the generation of stable ribosome-nascent chain complexes, the translation mixture

was incubated with 1 µl of 50 mM cycloheximide for 5 minutes on ice to terminate

translation without the release of nascent chains. Alternatively, in cases where the

release of nascent chains from the ribosome was necessary, 1.5 µl of 20 mM puromycin

and 0.5 µl of 250 mM EDTA were added, and the sample was incubated for 10 minutes

at 30 °C. The membrane fraction was recovered by centrifugation at ∼16,000g for 10

seconds. The resulting pellet was washed twice in 20 µl of KHM buffer and then

resuspended in 20 µl of KHM buffer before cross-linking reactions.


64

2.7 Isolation of ribosome-nascent chain complexes

For the synthesis of nascent chains in the absence of membranes, in vitro translations

were carried out as described above without the addition of semi-permeabilised cells.

The translation was halted by incubation with cycloheximide at the final concentration

of 2 mM for 5 minutes on ice. Isolation of the ribosome-nascent chain complexes was

achieved by layering the sample over 3 volumes of high salt/sucrose cushion (250 mM

sucrose, 500 mM KOAc, 5 mM Mg(OAc)2, 50 mM HEPES.KOH pH 7.9) and

centrifugation at 213,000g (70,000 rpm in a TLA120.2 rotor using a Beckmann Optima

TLX benchtop ultracentrifuge) for 20 minutes at 4 °C. The supernatant was discarded

and the pellet was resuspended in low salt/sucrose buffer (250 mM sucrose, 100 mM

KOAc, 5 mM Mg(OAc)2, 50 mM HEPES.KOH pH 7.9) for modification reactions.

2.8 Cross-linking and modifications of nascent chains with sulphydryl specific

reagents

All cross-linking and modification reagents were prepared in dimethylsulphoxide

(DMSO) as 20 mM stock solutions. For cross-linking, the membrane fraction

resuspended in KHM was incubated with the cross-linking reagent, BMH, at a final

concentration of 1 mM for 10 minutes at 30 °C. Similarly, for modification reactions,

sulphydryl specific reagents, AMS and QSY, were added to a final concentration of 1

mM and the sample was incubated for 10 minutes at 30 °C. β-mercaptoethanol was then

added to a final concentration of 5 mM to quench the cross-linking or modification

reaction. For control experiments, DMSO (5 % v/v) was added instead of the sulphydryl

specific reagent. RNase A (250 µg/ml) was added to all the samples and they were

incubated at 37 °C for 5 minutes in order to destroy any remaining peptidyl tRNA

species and the samples were then prepared for SDS-PAGE or subjected to

immunoprecipitation.

2.9 Solubilisation of ribosome-nascent chain complexes in C12E8

In vitro translations were carried out in the presence of semi-permeabilised cells and

halted either with cycloheximide or puromycin as described in section 2.6. The

membrane fraction was isolated by centrifugation through 3 volumes of high


65

salt/sucrose cushion at 130,000g for 10 minutes at 4 °C and resuspended in low

salt/sucrose buffer. C12E8 detergent was added to a final concentration of 1% (w/v) and

the sample was incubated on ice for 10 minutes. Ribosomal subunits and associated

nascent chains were recovered by centrifugation through a high salt/sucrose cushion

containing 0.1% (w/v) C12E8 at 213,000g for 20 minutes at 4 °C. The pelleted material

was solubilised in SDS sample buffer for analysis on SDS PAGE. The products in the

supernatant were subjected to trichloroacetic acid (TCA) precipitation. Hence, an equal

volume of a mixture of 20 % TCA (w/v) / 50 % acetone (v/v) was added to the

supernatant and incubated for 15 minutes on ice. The sample was spun at ∼16,000g for

20 minutes and the pellet was washed with 1 ml of 50 % acetone. The sample was spun

at ∼16,000g for an additional 20 minutes and the pellet was dried at 37 °C in a heat

block. The pellet was resuspended in 50 µl of SDS sample buffer and prepared for

loading onto an SDS-polyacrylamide gel (section 2.12).

2.10 Immunoprecipitation

After cross-linking or modification reaction, SDS was added to a final concentration of

1 % v/v and samples incubated at 37 °C for 30 minutes. At least 4 volumes of Triton IP

buffer (10 mM Tris-HCl pH 7.6, 140 mM NaCl, 1 mM EDTA, 1 % Triton X-100) was

added, together with 0.2 mg/ml protease inhibitor phenylmethyl sulphonyl fluoride, 1

mM methionine and 10 µl of a pansorbin suspension. The samples were left at 4 °C for

30 minutes on a rolling platform to preclear, spun at ∼16,000g for 20 minutes at 4 °C

and the supernatant transferred to fresh tubes. The supernatant was incubated with the

appropriate anti-serum (1 µl) and Protein A Sepharose (20 µl) at 4 °C overnight on a

rolling platform. The beads were spun down for 10 seconds at ∼16,000g and the

supernatant was discarded. The beads were washed three times by resuspending them in

IP buffer (800 µl) and spinning them down again. The samples were then prepared for

loading onto an SDS- polyacrylamide gel as described in section 2.12.

2.11 Endoglycosidase H digestion

Endoglycosidase H (Endo H) removes asparagine-linked oligosaccharide units by

cleaving between the two N-acetylglucosamine (GlcNAc) residues. Following washing

in IP buffer, immunoprecipitated samples on Protein A beads were denatured in Endo H


66

denaturing buffer (0.5% SDS, 1% β-mercaptoethanol) for 30 minutes at 37 °C. 3 µl of

0.5M sodium citrate (pH 5.5) buffer, 0.5 µl of 100 mM phenylmethyl sulphonyl

fluoride, 5.5 µl of H2O and 500 U Endo H were added and the samples were incubated

for an additional 30 minutes at 37 °C. The samples were then solubilised in an equal

volume of 2x concentration SDS sample buffer and resolved by SDS PAGE.

2.12 SDS PAGE and sample analysis

The samples were first denatured in 50 µl of SDS sample buffer (0.1 M Tris-HCl pH

6.8, 5 mM EDTA, 0.5 M sucrose, 0.5 % L-methionine, 0.01 % bromophenol blue, 0.5

M DTT, 20% SDS) by incubation at 37 °C for 30 minutes. Cross-linking samples were

loaded onto a denaturing 14% polyacrylamide gel, while samples from modification

reactions with AMS or QSY were loaded onto a denaturing 18% polyacrylamide gel.

These samples were run in the presence of SDS and resolved at 150 V. The gels were

then fixed in acetic acid: methanol: water solution (1:2:7 v/v) for at least 5 minutes

before they were dried. They were then exposed to a phosphoimaging plate for three to

seven days and the results analysed on a Fuji BAStation phosphoimager using the

AIDA software. Where specified, quantification of the products was also performed

using the AIDA software.

Chapter 3 – The use of a site-specific cross-linking approach

67

CHAPTER 3 Results

The use of a site-specific cross-

linking approach to examine opsin

integration

Chapter 3 – The use of site-specific cross-linking approach

68

3.1 Introduction

An essential step during membrane protein biosynthesis at the ER is the integration of

its one or more transmembrane (TM) domains. This process involves the lateral

movement of the TM domain from the proteinaceous environment of the Sec61

translocon to the phospholipid bilayer of the ER membrane. Although the crystal

structure of an archeal Sec61 complex has been elucidated (Van den Berg et al., 2004),

the mechanism by which integration occurs is unknown. The insertion of a polytopic

membrane protein is a particularly complex process since several TM domains have to

be correctly integrated in order to enable the proper assembly of a functional

polypeptide. The aim of this study is to use the seven transmembrane domain protein,

opsin, as a model to examine the biosynthesis of polytopic membrane proteins and to

establish how the integration of multiple TM domains is co-ordinated.

Opsin has several characteristics that makes it well-suited to the study of polytopic

membrane protein biogenesis: 1) it belongs to a well-characterised family of seven

transmembrane domain receptor proteins; 2) extensive structural data is available from a

crystal structure (Palczewski et al., 2000); 3) a good monoclonal antibody to its N-

terminal region is available; 4) the two N-glycosylation sites near its N-terminus serve

as ideal marker for the authentic targeting and insertion of nascent opsin chains into the

ER membrane (c.f. Laird & High, 1997).

In this study, the principal approach employed to examine opsin integration is to

analyse the molecular environment of each TM domain at different stages of opsin

biosynthesis by using a site-specific cross-linking approach. To this end, all the natural

cysteines in the wild type opsin sequence were replaced (Meacock et al., 2002), and a

single cysteine residue was introduced into the particular TM domain of interest. This

approach ensures that the cross-linking reaction can only occur from the single cysteine

residue introduced into the nascent chain. In order to mimic specific stages of opsin

biosynthesis, nascent chains of specific lengths were generated by the translation of

truncated mRNAs. Since these mRNAs lack a stop codon, the nascent chains are not

released from the ribosome, but remain lodged at the ER translocon, creating artificial

‘integration intermediates’ (Gilmore et al., 1991). The addition of the homo-

bifunctional thiol-reactive reagent, bismaleimidohexane (BMH), at this stage allows the


69

single cysteine probe within the nascent chain to be cross-linked to any available

cysteine residues present in adjacent proteins. By performing the cross-linking reaction

with integration intermediates of increasing nascent chain lengths, the molecular

environment of a specific TM domain could be examined at different stages of opsin

biosynthesis.

3.2 Optimisation of the experimental system

The analysis of opsin biogenesis using cross-linking techniques is well-established

(Laird & High, 1997) and has been extensively used to examine the environment of

TM1 and TM2 of opsin (Meacock et al., 2002). In this approach, an integration

intermediate of a particular chain length is taken to reflect a particular stage of

membrane integration (Thrift et al., 1991). However, previous studies of opsin have

shown that the population of nascent chains generated by this approach may not be

homogeneous, thus, the presence of nascent chains of differing lengths within a single

translation reaction can result in complex cross-linking patterns that may be difficult to

interpret unambiguously (Meacock et al., 2002).

One obvious factor that might contribute to the heterogeneity of the nascent chain

population is ribosome stacking. Multiple ribosomes often translate a single mRNA and

under normal circumstances, this leads to multiple copies of the full length protein.

However, in the case of the truncated mRNAs that are used in vitro, the ribosomes are

not released from the mRNA and may stack up at its 3’ end generating different length

nascent opsin chains, some of which lack the C-terminal region (Fig. 3.1). For

meaningful cross-linking experiments, it is imperative that cross-linking adducts formed

from authentic integration intermediates can be distinguished from any adducts that

might be formed from such shorter nascent chains. For this purpose, a haemagglutinin

(HA) epitope tag was introduced at the C-terminus of each integration intermediate. By

performing immunoprecipitation with an α-HA antiserum, only nascent chains with an

intact C-terminus (i.e. nascent chains of the correct length) will be recovered, thus

allowing ‘authentic’ cross-linking adducts to be identified (Fig. 3.1).

In order to establish whether the C-terminal HA-tagging of integration intermediates

allows the efficient selection of authentic nascent chains, two opsin constructs of


70

different lengths were used. OP164[cys56] has a cysteine residue at position 56 with the

first 155 residues of opsin followed by the 9 residues which make up the HA tag at the

C-terminus (total 164 residues). The longer construct, OP357[cys56], also has a

cysteine residue at position 56 with the complete 348 residues of the opsin coding

region plus a 9-residue HA tag (total 357 residues).

Figure 3. 1 Rationale for HA tagging of nascent opsin chains. Opsin chains of different lengths may arise as a consequence of ribosome stacking where several ribosomes are synthesizing polypeptide chains from a single truncated mRNA chain (in blue). These shorter chains will lack the C-terminal HA tag (in green) but will still be immunoprecipitated by the α-opsin monoclonal antibody which recognises the N-terminus of the polypeptide. Adducts with these shorter chains will be recognised by sera specific for subunits of the ER translocon. However, only authentic opsin chains with an intact C-terminal HA tag will be immunoprecipitated by the α-opsin, α-Sec61 subunit and α-HA antisera.

mRNA representing each integration intermediate was translated in vitro in the presence

of semi-permeabilised mammalian cells and incubated with either BMH or a solvent

(DMSO) only control. After denaturation with 1% SDS, the nascent chains were

immunoprecipitated with the α-opsin antisera which recognises the N-terminus of opsin,

and the α-HA antisera which recognizes the C-terminal HA tag (Fig. 3.2).

The proper integration of the nascent opsin chains into the ER membrane of the semi-

permeabilised cells was confirmed by the presence of doubly N-glycosylated chains

(Fig. 3.2, denoted by (ii)). The lower product observed in all the samples represents the

fraction of opsin chains that are not glycosylated (Fig. 3.2, (i)). Several additional


71

products were also present following immunoprecipitation with the α-opsin antisera, but

were absent after immunoprecipitation with the α-HA antisera (Fig. 3.2, c.f. lanes 1 and

2 to 3 and 4, and lanes 5 and 6 to 7 and 8, denoted by a bracket). The lack of the HA tag

confirms that these polypeptide chains were truncated at their C-termini and may have

been generated by ribosome stacking or by the degradation of longer opsin chains.

These truncated products were especially prevalent for the longer polypeptide,

OP357[cys56] (Fig. 3.2, c.f. lanes 1 and 2 to 5 and 6, (])), thus highlighting the

importance of C-terminal HA-tagging particularly for long nascent chains.

In the presence of BMH, two distinct cross-linking adducts were observed for

OP164[cys56] while one adduct was seen for OP357[cys56] (Fig. 3.2, lanes 2, 4, 6 and

8, (X)). These adducts were immunoprecipitated with the α-HA antisera (Fig. 3.2, lanes

4 and 8, (X)), indicating that the cross-linking products were formed with nascent

chains of the appropriate lengths. In this way, the inclusion of the HA tag at the C-

terminus of the integration intermediate allows authentic cross-linking adducts to be

distinguished from any other adducts that may be formed with any shorter nascent

chains present in the translation reaction.

3.3 Cross-linking adducts are formed with glycosylated opsin chains

In order to examine the molecular environment of opsin TM domains during nascent

chain biogenesis by cross-linking, it is important that cross-linking products are formed

with nascent chains that are properly integrated into the membrane. N-glycosylation of

the N-terminal region of opsin serves as a useful marker for efficient nascent chain

integration into the ER membrane. One way to ascertain that the BMH-dependent cross-

linking adducts are formed with glycosylated nascent opsin chains is by treating the

samples with endoglycosidase H. Endoglycosidase H cleaves the glycan group attached

to the opsin chain between the two N-acetylglucosamine (GlcNAc) residues, resulting

in faster migration in SDS-PAGE due to a reduction in molecular weight.

The construct, OP109[cys56], has a single cysteine residue located within TM1 of opsin

and a chain length of 109 residues, including the HA tag. mRNA encoding

OP109[cys56] was translated in the presence of semi-permeabilised cells, treated with

either BMH or DMSO and subjected to immunoprecipitation using α-opsin, α-HA, α-


72

Sec61α and α-Sec61β antisera. A duplicate set of samples were additionally treated with

endoglycosidase H.

Figure 3. 2 Immunoprecipitation with α-HA antisera allows selection of authentic opsin chains. mRNA representing OP164[cys56] and OP357[cys56] were translated in vitro in a rabbit reticulocyte translation system in the presence of semi-permeabilised mammalian cells and radiolabelled methionine. The membrane fraction was isolated by centrifugation and treated with either BMH or DMSO (solvent control). The cross-linking reaction was then quenched with β-mercaptoethanol. The samples were denatured in 1% SDS and immunoprecipitations using α-opsin and α-HA antisera were carried out. Uncross-linked doubly-glycosylated opsin chains resulting from both constructs are denoted by (ii), while uncross-linked non-glycosylated opsin chains are indicated with (i). Truncated opsin chains which lack an intact C-terminal HA tag are present in immunoprecipitations with the α-opsin antibody (denoted by brackets (])), but were absent in immunoprecipitations with the α-HA antibody. BMH-dependent cross-linking adducts were also observed for both integration intermediates and are marked by a cross (X).


73

Figure 3. 3 Cross-linking products are formed with glycosylated opsin chains. OP109[cys56] was synthesised in a rabbit reticulocyte translation system in the presence of digitonin-permeabilised mammalian cells. The membrane fraction was incubated either with BMH (+) or mock-treated with solvent only (-). Denaturing immunoprecipitation was then carried out using α-opsin, α-HA, α-Sec61α and α-Sec61β antisera. Duplicate samples were treated with endoglycosidase H (+ EndoH) after immunoprecipitation. Distinct cross-linking adducts obtained in the presence of BMH were marked with a cross (X). Products that were identified as adducts with Sec61α and Sec61β are indicated with ‘α’ and ‘β’ respectively. Other symbols are as previously defined in the legend to Figure 3.2.


74

Both glycosylated and non-glycosylated OP109[cys56] nascent chains were observed

for samples which were not treated with endoglycosidase H (Fig. 3.3, lanes 1-3, denoted

by (ii) and (i) respectively). As before, immunoprecipitation with the α-HA antisera

identified authentic nascent chains and their corresponding BMH-dependent cross-

linking adducts (Fig. 3.3, lane 3, (i), (ii), (X)). The adducts were identified by

immunoprecipitation as nascent chains cross-linked to Sec61α and Sec61β (Fig. 3.3,

lanes 4 and 5, (α), (β)). Treatment of the samples with endoglycosidase H resulted in the

collapse of uncross-linked glycosylated opsin chains to a single lower molecular weight

band which represented the de-glycosylated nascent chains (Fig. 3.3, lanes 6-8, (i)).

The OP109[cys56] adducts with Sec61α and Sec61β also migrated faster following

treatment with endoglycosidase H, as shown by the shift of these products upon SDS-

PAGE, indicating a reduction in molecular weight (Fig. 3.3, c.f. lanes 4 and 5 to 9 and

10, (α), (β)). The sensitivity of the adducts to endoglycosidase H treatment indicated

that both Sec61α and Sec61β were cross-linked to opsin chains which were N-

glycosylated. Thus, these cross-linking products were formed with nascent chains which

have been correctly targeted and integrated into the ER membrane.

3.4 Cross-linking adduct formation is cysteine-dependent

The ‘site-specificity’ of the cross-linking reaction is pivotal to the use of cross-linking

techniques in the analysis of the environment of different TM domains during their

integration into the ER membrane. The use of a homo-bifunctional, cysteine reactive

cross-linking reagent such as BMH ensures that adducts are formed only from the target

cysteine probe present in the TM domain. However, previous studies have shown that

spontaneous non-specific cross-linking reactions may still occur in the absence of a

cross-linking reagent (Oliver et al., 1996). In order to establish that the adducts

observed in the previous experiments were cysteine dependent, cross-linking was

performed with opsin constructs which lack a cysteine probe.

OP164[cys-null] and OP204[cys-null] are two integration intermediates with chain

lengths 164 and 204 respectively including the C-terminal HA tag, which do not contain

any cysteine probes. In vitro translation and the cross-linking reaction were performed

as before and the products were immunoprecipitated using α-opsin, α-HA, α-Sec61α


75

and α-Sec61β antisera. No distinct cross-linking adducts were observed for both

integration intermediates, indicating that BMH dependent cross-linking reaction

requires the presence of a cysteine probe in the nascent chains (Fig. 3.4, lanes 1-3 and 6-

8).

However, immunoprecipitation with the α-Sec61α antibody did result in two products of

∼40 kDa and ∼80 kDa (Fig. 3.4, lanes 4 and 9, (•)). These products were not recognized

by the α-opsin or α-HA antisera, indicating that they do not contain any opsin chains

(Fig. 3.4, c.f lanes 2 and 3 to 4, c.f. lanes 7 and 8 to 9). On the basis of size alone, the 40

kDa band may be due to endogenous, radiolabelled Sec61α molecules. As BMH was

present in the sample, the larger band of ∼80 kDa may consequently be due to two

molecules of endogenous Sec61α cross-linked together.

In order to determine if these products were due to radiolabelled Sec61α molecules, the

translation reaction was repeated without the addition of any exogenous mRNA

template and BMH cross-linking was performed (Fig. 3.5). Immunoprecipitation with

the α-Sec61α antibody gave a single product of ∼40 kDa in the absence of BMH (Fig.

3.5, lane 3, (•)), while two species of ∼40 kDa and ∼80 kDa were observed in the

presence of BMH, reminiscent of the products seen in the previous experiment (c.f. Fig.

3.5, lane 4, to Fig. 3.4, lanes 4 and 9, (•)). This confirmed that the ∼40 kDa band was an

endogenous, radiolabelled Sec61α monomer while the ∼80 kDa band was most likely

due to two Sec61α molecules cross-linked together. Immunoprecipitation using the α-

opsin and α-Sec61β antisera did not give any products, indicating that no endogenous

radiolabelled opsin or Sec61β molecules were present (Fig. 3.5, lanes 1, 2, 5 and 6).

The radiolabelled Sec61α species seen in Figures 3.4 and 3.5 were generated when

endogenous Sec61α mRNA associated with the semi-intact cells was translated in the

presence of radiolabelled methionine. Although the semi-permeabilised cells used in

these assays were treated with micrococcal nuclease, the nuclease treatment was clearly

not completely effective in removing all endogenous mRNA chains, thus allowing

traces of other molecules to be radiolabelled during the translation reaction. The

presence of these radiolabelled Sec61α species in the sample further highlighted the

need for strict criteria in distinguishing true cross-linking adducts from other species.

Hence, only if a cross-linking product is present in both the α-HA and α-opsin samples,


76

will it be considered as a bona fide adduct formed from an authentic opsin integration

intermediate.

Figure 3. 4 Formation of BMH cross-linking adducts is cysteine dependent. mRNA representing OP164[cys-null] and OP204[cys-null] was translated in the presence of semi-permeabilised cells and the membrane fraction was treated with either BMH (+) or DMSO (-). As before, immunoprecipitations with α-opsin, α-HA, α-Sec61α and α-Sec61β antisera were carried out. No distinct BMH-dependent cross-linking adducts were observed. However, immunoprecipitation with the α-Sec61α antibody gave two faint products (denoted with (•)).


77

Figure 3. 5 Radiolabelled endogenous Sec61α molecules were generated during translation. In vitro translation was performed without the addition of exogenous mRNA templates in the presence of radiolabelled methionine and digitonin-permeabilised cells. The sample was then subjected to denaturing immunoprecipitations using α-opsin, α- Sec61α and α-Sec61β specific antisera. Sec61α specific products obtained by immunoprecipitation with the α-Sec61α antibody are indicated with (•).


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3.5 Summary

For cross-linking to be successfully exploited in the study of opsin integration, it is

necessary to have specific controls to allow an unambiguous interpretation of the

resulting data. Crucial factors which have been addressed are: 1) the cross-linking

reaction is cysteine dependent, i.e. the reaction is site-specific; 2) the cross-linking

adducts are formed with glycosylated (i.e. properly integrated) opsin chains; 3) adducts

formed with authentic opsin chains of the correct length may be distinguished from

adducts formed with shorter, truncated nascent chains by the addition of a C-terminal

HA tag. The inclusion of a C-terminal epitope tag is particularly important for long

chain lengths.

Conclusion

Having established the validity of this experimental system, I now set out to exploit it in

order to analyse the molecular environment of specific regions of opsin, in particular its

TM domains, during membrane integration.

Chapter 4 – The analysis of TM1, TM2 and TM3 of opsin

79

CHAPTER 4 Results

The membrane integration of the

N-terminal region of opsin:

TM1 to TM3


80

4.1 Introduction

This study aims to analyse the molecular environment of each of the seven TM domains

of opsin during its membrane integration in order to gain insights into the mechanism of

polytopic membrane protein biogenesis at the ER. In the previous chapter, the cysteine

mediated site-specific cross-linking approach employed during this investigation was

validated. Thus, unique cysteine residues can be placed at strategic locations within the

nascent chain and their environment probed using the homo-bifunctional, sulphydryl-

specific cross-linking reagent, BMH. A second feature of this approach is the use of

defined integration intermediates that can be generated in vitro. It had previously been

suggested that the presence of incomplete intermediates, generated either by ribosome

stacking or degradation of longer opsin chains, may complicate the interpretation of

such studies (Meacock et al., 2002). I showed that the addition of a C-terminal HA tag

to opsin integration intermediates allowed authentic chains and their cross-linking

products to be distinguished from other ‘spurious’ adducts. For this reason, all

integration intermediates generated in this study included the 9 residue HA epitope tag

at the C-terminus.

Previous studies of opsin fragments suggest it is comprised of two or more independent

folding domains (Ridge et al., 1995). The aim of the work presented in this chapter was

to investigate the molecular environment of the N-terminal region of opsin, comprising

TM1 to TM3 during membrane integration of the nascent polypeptide. Thus, a single

cysteine residue was introduced into a particular TM domain of a cysteine-null opsin

mutant to allow site-specific cross-linking from the target cysteine to ER components in

close proximity. In order to follow the location of a particular TM domain during its

integration, a range of integration intermediates of increasing chain lengths was

generated to represent different stages of opsin biosynthesis (as depicted in Figure 4.1).


81

Figure 4. 1 A diagrammatic representation of artificial opsin integration intermediates generated for site-specific cross-linking analysis of distinct TM domains. All cysteine residues of wild type opsin were replaced with glycine (Meacock et al., 2002) and a single cysteine probe was introduced into TM1, TM2 or TM3 (for simplicity, the cysteine residue is not shown in this diagram). Transmembrane domains are represented with a dashed line while the C-terminal HA tag is indicated in grey. The numbering shows the location of the TM domains (derived from the crystal structure, see Palczewski et al., 2000) in the context of the hypothetical topologies of the various integration intermediates. The two N-linked glycosylation sites at residues 2 and 15 are indicated with a ‘Y’.


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4.2 TM1 is adjacent to discrete sets of ER components during its integration

Whilst the integration of opsin TM1 has been previously characterised by a cysteine-

mediated cross-linking study, the heterogeneity of the nascent chain population present

in this study complicated the interpretation of the data (Meacock et al., 2002). In order

to obtain a more accurate analysis of TM1 integration, nascent opsin chains of the same

lengths as those previously examined but including the HA epitope tag at their C-

terminus, were generated. These integration intermediates possess a single cysteine at

residue 56 within TM1 of opsin which acts as the sole target for the subsequent cross-

linking reaction.

mRNA transcripts representing integration intermediates of OP[cys56] were translated

in vitro in the presence of digitonin-permeabilised mammalian HT1080 cells. The

membrane fraction was isolated by centrifugation and was then either treated with the

sulphydryl-specific cross-linking reagent, BMH, or mock-treated with solvent alone

(DMSO). Following quenching of any unreacted maleimides and the denaturation of the

samples using 1% SDS, specific products were recovered by immunoprecipitation using

α-opsin, α-HA, α-Sec61α and α-Sec61β antisera. As previously observed (see Chapter

3), there was clear heterogeneity in the population of nascent chains that were

membrane-integrated (Fig. 4.2). This was largely restricted to longer intermediates, i.e.

OP150[cys56] to OP259[cys56], resulting in several products that were recognised by

the α-opsin but not the α-HA antibody (Fig. 4.2, c.f. lanes 17 and 18, 22 and 23, 27 and

28, 32 and 33, see *). In some cases, these incomplete intermediates gave adducts to ER

translocon components, but such adducts could be distinguished from those formed with

authentic integration intermediates since they were not immunoprecipitated with the α-

HA antibody (Fig. 4.2, c.f. adducts to Sec61α in lanes 13 and 14, 28 and 29).

In contrast, a number of cross-linking products were found to be efficiently

immunoprecipitated with both the α-opsin and α-HA antisera, indicating these adducts

resulted from authentic integration intermediates cross-linked to ER components (Fig.

4.2, lanes 2 and 3, 7 and 8, 12 and 13, 17 and 18, 22 and 23, 27 and 28, 32 and 33).

Several of these products were identified as adducts with either subunits of the Sec61

complex, or a previously defined novel 10 kDa protein PAT-10 (a protein associated


83

with the translocon of ~10 kDa) (Meacock et al., 2002) (Fig. 4.2, lanes 2-5, 7-10, 12-13,

17-19, 22-24, 27-28, 32-33, products denoted by α, β and P respectively).

Figure 4. 2 BMH-mediated cross-linking of OP[cys56] integration intermediates of increasing chain length to ER components. Truncated mRNA chains representing the integration intermediates indicated were translated in rabbit reticulocyte lysate supplemented with digitonin-permeabilised mammalian cells. Membrane-integrated products were isolated by centrifugation and incubated with either BMH (+) or DMSO (-). The samples were quenched with β-mercaptoethanol, denatured in 1% SDS and specific products were recovered by immunoprecipitations using α-opsin, α-HA, α-Sec61α and α-Sec61β antisera. Uncross-linked doubly-glycosylated and non-glycosylated opsin chains are denoted with (ii) and (i) respectively, while truncated opsin chains which lack an intact C-terminus are marked with asterisks. Distinct adducts to Sec61α, Sec61β and PAT-10 are indicated with ‘α’, ‘β’ and ‘P’ respectively.


84

It was striking that cross-linking to Sec61β was seen only with short opsin chains, OP96

and OP109 (Fig. 4.2, β), while cross-linking to PAT-10 was apparent with OP130 and

beyond (Fig. 4.2, P). Whilst the exact identity of PAT-10 remains unclear, it has been

extensively characterised during the previous analysis of TM1 integration (Meacock et

al., 2002).

Surprisingly, the cross-linking of opsin TM1 to Sec61α displayed a previously

unrecognised level of complexity (c.f. Meacock et al., 2002). Thus, TM1 formed

distinct adducts to Sec61α with the shorter opsin chains, OP96 and OP109, but adduct

formation was barely detectable when the nascent chain length was increased to 130

residues (Fig. 4.2, lanes 4, 9 and 14). Remarkably, extending the nascent chain further

to 150 and 164 residues resulted in a second association with Sec61α that was then

diminished again upon further chain extension to OP204 and OP259 (Fig. 4.2, lanes 19,

24, 29 and 34, α, see also Fig. 4.4 for quantification). These results indicated that TM1

experiences at least two distinct Sec61 mediated environments during opsin

biosynthesis; one in which TM1 may be cross-linked to both Sec61α and Sec61β, and

another where TM1 is adjacent to Sec61α and PAT-10. Taken together, these data

suggest TM1 engages the translocon as soon as it emerges from the ribosome, moves

away upon chain extension, but then transiently re-associates with the translocon in an

environment close to PAT-10.

4.3 TM1 environment is influenced by subsequent TM domains

The alteration in the environment of TM1 observed by cross-linking of different

integration intermediates is most likely a consequence of TM1 relocation during nascent

chain extension. If so, then the environment of TM1 may alter either as a result of

‘active displacement’ by the subsequent TM domains that are synthesised as the nascent

opsin chain gets longer, or alternatively, the change may simply result from having a

longer polypeptide ‘tether’ to the ribosome thereby allowing TM1 to relocate to a new

environment. In order to distinguish between these two possibilities and determine

whether the subsequent TM domains have an influence on TM1 relocation, TM2 to

TM7 of opsin were replaced with a hydrophilic region of the secretory protein

preprolactin to produce the OPTM1PPL[cys56] polypeptide (see Fig. 4.3A). The

environment of TM1 during nascent chain extension was then analysed using


85

integration intermediates of OPTM1PPL[cys56] with lengths identical to those

previously used to analyse the opsin nascent chain (c.f. Fig. 4.2).

The site-specific cross-linking products of a range of OPTM1PPL[cys56] integration

intermediates were analysed by immunoprecipitation as previously described. Authentic

adducts, i.e. recognised by both α-opsin and α-HA antisera, were observed between the

short OPTM1PPL109[cys56] nascent chain and both Sec61α and Sec61β (Fig. 4.3B,

lanes 2-5). These adducts were found to persist at longer chain lengths of 130, 150, 164

and 204 residues (Fig. 4.3B, lanes 7-10, 12-15, 17-20, 22-25, α and β) and this

prolonged association of TM1 with the Sec61 complex when present in

OPTM1PPL[cys56] was clearly distinct from its previous behaviour (c.f. Fig. 4.2).

Whilst immunoprecipition using antisera recognising Sec61α and Sec61β also

recovered products with the OPTM1PPL259 intermediate, these adducts were not

significantly immunoprecipitated with the α-HA antibody, indicating that they were

adducts to shorter nascent chains lacking the C-terminal HA epitope (Fig. 4.3B, c.f.

lanes 29-30). It was also apparent that TM1 of the OPTM1PPL construct was cross-

linked to PAT-10 from a chain length of 130 residues through to 259 residues, although

the efficiency of cross-linking was substantially reduced in comparison to the opsin

chains (c.f. Fig. 4.2 and 4.3B, lanes 8, 13, 18, 23, 28, P). Adducts with subunits of the

Sec61 complex and PAT-10 were not observed when OPTM1PPL integration

intermediates were released with puromycin prior to BMH cross-linking (data not

shown).

In order to allow a better comparison between the behaviour of TM1 in the context of

both OP[cys56] and OPTM1PPL[cys56], the relative cross-linking efficiency of Sec61α

was determined in both cases (Fig. 4.4). This reaffirmed that, whilst TM1 experiences

two distinct Sec61 based environments when present in the opsin chain, this periodicity

is not observed when it is present in OPTM1PPL. In OPTM1PPL, TM1 cross-linking to

Sec61α remained fairly constant until the nascent chain was 259 residues long. Taken

together, with the distinct difference in Sec61β adduct formation between the two

constructs, these results suggest that the synthesis of the subsequent TM domains (TM2

to TM7) influences TM1 relocation from the Sec61 complex and that the absence of

these TM domains causes a delay in its exit from the ER translocon.


86

Figure 4. 3 A) A schematic representation of the OPTM1PPL[cys56] polypeptide. The first 70 residues of opsin including TM1 (in black) were fused to a region of the secretory protein, preprolactin (PPL, residues 31-229, in grey) to form OPTM1PPL. A single cysteine probe was present at position 56 within TM1 (indicated by a white star). B) BMH cross-linking of OPTM1PPL[cys56] integration intermediates of to translocon associated components. Integration intermediates of different lengths were synthesised in a rabbit reticulocyte translation system in the presence of semi-permeabilised cells and treated as described in the legend to Figure 4.2. Uncross-linked doubly-glycosylated, non-glycosylated and truncated nascent opsin chains are denoted with (ii), (i) and asterisks respectively, while cross-linking adducts to Sec61α, Sec61β and PAT-10 are labelled ‘α’, ‘β’ and ‘P’ respectively.


87

Figure 4. 4 A plot of the relative efficiency of cross-linking to Sec61α versus the chain length of the integration intermediate. For relevant integration intermediates, quantification of the uncross-linked doubly-glycosylated opsin chains (e.g. Fig. 4.3B, lane 1, (ii)) and its adducts with Sec61α (e.g. Fig. 4.3B, lane 2, α) uses the products immunoprecipitated with the α-opsin antibody and was carried out using the AIDA software (see appendix for raw data). The fraction of nascent chains cross-linked to Sec61α was initially calculated by dividing the amount of the Sec61α adduct by the total amount of glycosylated opsin chains present in the cross-linking reaction. The integration intermediate for which the highest fraction of nascent chain was cross-linked to Sec61α was then set as the nominal value of 1.0 and other levels of adduct formation were expressed relative to the highest level. These values (y-axis) were then plotted against the length of the nascent chain (x-axis). In the case of normal opsin (represented by solid line), OP96 was set as the reference point, while in the case of OPTM1PPL (represented by dashed line), OPTM1PPL164 was set as the reference point.


88

4.4 TM2 has exited the translocon in the OP204 integration intermediate

An extensive analysis of TM2 integration has been carried out in a previous study of

opsin biosynthesis (Meacock et al., 2002), therefore less emphasis was placed on

investigating the molecular environment of TM2 in this study. However, the alteration

of the behaviour of TM1 when examined in the context of the OPTM1PPL raises the

question of the relative position of the other TM domains with respect to TM1 when

present in the normal opsin construct with multiple TM domains. The environment of

integration intermediates with cys89 located within TM2 of three different chain

lengths, OP140, OP164 and OP204, was therefore analysed to examine the location of

TM2.

OP140[cys89], OP164[cys89] and OP204[cys89] chains were treated with BMH and

recovered by immunoprecipitation as previously described. Adducts with Sec61α and

Sec61β were observed with OP140 and OP164 opsin chains (Fig. 4.5, lanes 2-5, 7-10, α

and β), but when the nascent chain is extended to 204 residues, no distinct adducts to

translocon components were seen (Fig. 4.5, lanes 12-15). These results indicate that

TM2 is engaged with the translocon when the nascent chain is 140 or 164 residues long,

but TM2 is presumed to have exited the translocon when the nascent chain is 204

residues.


89

Figure 4. 5 BMH cross-linking of integration intermediates with cys89. OP140[cys89], OP164[cys89] and OP204[cys89] were synthesised, treated with BMH and recovered by immunoprecipitation as described in the legend of Figure 4.2. Symbols used are as previously defined. This figure was provided by Samuel Crawshaw.


90

4.5 TM3 is associated with the ER translocon in the OP164 integration

intermediate

OP164 represents an important stage in opsin biosynthesis where changes in the

environment of TM1 seem to occur, where TM1 was seen to regain association with

Sec61α (Fig. 4.2). On the basis of chain length, one might expect opsin TM3 to have

exited the ribosome (see Fig. 4.1) in the OP164 integration intermediate and hence the

environment of TM3 was investigated at this point. As opsin TM3 is a long TM domain

which consists of residues 107 to 139 (Palczewski et al., 2000), its molecular

environment was examined using probes introduced at three different locations. Thus,

single cysteine probes were placed towards the extracellular end of TM3 (cys115), in

the middle of TM3 (cys124) or towards the intracellular end of TM3 (cys132).

Integration intermediates of OP164[cys115], OP164[cys124] and OP164[cys132] were

cross-linked to adjacent ER proteins using BMH and adducts identified by

immunoprecipitation. The resulting adducts were quite distinct for the three different

cysteine residues used, and whilst discrete adducts with both Sec61α and Sec61β were

observed using OP164[cys115] (Fig. 4.6, lanes 2-5, α and β), only an adduct to Sec61β

was seen with OP164[cys124] (Fig. 4.6, lanes 7-10, β). On the other hand, no

significant adducts to any of these Sec61 subunits were seen with OP164[cys132] (Fig.

4.6, lanes 12-15). It is quite possible that cys132 has not fully exited the ribosome at a

chain length of 164 residues and is therefore not sufficiently close to the ER translocon

for efficient cross-linking to occur. On the basis of adduct formation between cysteine

probes 115 and 124, and subunits of the Sec61 complex, I conclude that TM3 has begun

to engage the translocon at its N-terminal end when the opsin chain has reached 164

residues in length.


91

Figure 4. 6 BMH cross-linking of OP164 integration intermediates with cysteine probes in three different locations within TM3. mRNA chains representing OP164[cys115], OP164[cys124] and OP164[cys132] were translated in vitro in the presence of semi-permeabilised cells and treated as described in the legend of Figure 4.2. As before, uncross-linked doubly-glycosylated and non-glycosylated opsin chains are indicated with (ii) and (i) respectively, while truncated opsin chains lacking a C-terminal HA tag are denoted by asterisks. Distinct cross-linking adducts to Sec61α and Sec61β are indicated with ‘α’ and ‘β’.


92

4.6 TM3 exits the translocon upon chain extension

Since cys115 of TM3 gave clear adducts to both the Sec61α and Sec61β subunits in

OP164, the effect of altering chain length upon the cross-linking partners of TM3 was

investigated by using integration intermediates of different lengths, all containing a

cysteine probe at position 115. In this case, integration intermediates ranging from

OP150[cys115] to OP357[cys115] were subjected to BMH mediated cross-linking and

the resulting products identified by immunoprecipitation.

At a chain length of 150 residues, the shortest analysed in this case, adducts with the

Sec61β subunit (Fig. 4.7, lanes 2-5, β) and a ∼21 kDa component that was similar to a

previously identified ribosomal protein were observed (Fig. 4.7, lanes 2-3, R) (Laird &

High, 1997). Thus, at this stage, opsin TM3 appears to be in the process of leaving the

ribosomal exit site and engaging the ER translocon. Distinct adducts with both Sec61α

and Sec61β were seen for the longer integration intermediates, OP164[cys115] and

OP174[cys115], indicating that cys115 of TM3 has now fully engaged the ER

translocon (Fig. 4.7, lanes 7-10, 12-15, α and β). A higher molecular weight product

representing species containing HA-tagged opsin chains and both the α and β subunits

of the Sec61 complex were also observed with OP164[cys115] and OP174[cys115]

(Fig. 4.7, lanes 9-10, 14-15, αβ).

On the other hand, no authentic adducts to subunits of the Sec61 complex were seen for

nascent chains longer than 174 residues (Fig. 4.7, lanes 17-21, 23-27, 29-33, 35-39, 41-

45). In some cases, endogenous radiolabelled Sec61α molecules were

immunoprecipitated with the Sec61α antibody (Fig. 4.7, •, see also Chapter 3.4), while

products immunoprecipitated with α-Sec61β were adducts formed with incomplete

opsin chains which lacked the C-terminal HA tag (Fig. 4.7, c.f. 18 and 21, 24 and 27, 30

and 33, 36 and 39, 42 and 45, ). On this basis, I conclude that TM3 fully engages the

translocon from a nascent chain length of 164 residues and has exited the translocon

when the nascent chain is 204 residues long. Since the extension of the nascent chain

from 164 residues to 204 residues results in opsin TM4 (residues 151 to 173)

(Palczewski et al., 2000) being synthesised in full and predicted to have largely exited

the ribosome, TM3 is likely to have left the Sec61 complex when TM4 begins to engage

the translocon (see Chapter 5).


93

Figure 4. 7 BMH mediated cross-linking of integration intermediates containing TM3 specific cysteine probes to ER translocon components. Integration intermediates of the chain lengths indicated and containing cys115 probes were synthesised in a rabbit reticulocyte translation system in the presence of digitonin-permeabilised cells and treated with either BMH (+) or DMSO (-). Samples were denatured in 1% SDS and subjected to immunoprecipitation using α-opsin, α-HA, α-Sec61α, α-Sec61β and a non-related antisera (NS). Uncross-linked doubly- and non-glycosylated opsin chains are denoted with (ii) and (i) respectively. Endogenous, radiolabelled Sec61α molecules which were immunoprecipitated by the Sec61α antibody are marked a filled circle while Sec61β adducts to truncated opsin chains are denoted with an empty circle. Cross-linking products to Sec61α and Sec61β are indicated by ‘α’ and ‘β’ while nascent chains cross-linked to both Sec61α and Sec61β are marked with ‘αβ’. Adducts with a putative ∼21 kDa ribosomal protein are also indicated, see lanes 2 and 3, R (see also Laird and High, 1997).


94

4.7 TM3 relocation is independent of the presence of subsequent TM domains

TM3 appeared to be fully engaged with the Sec61 translocon when the opsin chain was

164 residues long and to have left the Sec61 complex by the time the nascent chain

reached 204 residues. As with the TM1 analysis in Section 4.3, the influence of

subsequent TM domains (i.e. TM4 to TM7) on the relocation of TM3 was investigated.

The region of opsin C-terminal of TM3, including TM4 to TM7, was replaced with a

hydrophilic region from the secretory protein, preprolactin, to give the OPTM1-

3PPL[cys115] construct (Fig. 4.8A). Integration intermediates of 164 and 204 residues,

comparable to those examined during the analysis of TM3, were generated.

The integration intermediates of this chimera, OPTM1-3PPL164[cys115] and OPTM1-

3PPL204[cys115], were then analysed by BMH mediated cross-linking as before (c.f.

section 4.2). OPTM1-3PPL164[cys115] gave a very similar cross-linking pattern to

OP164[cys115] displaying clear adducts with Sec61α and Sec61β (Fig. 4.8B, lanes 2-5,

c.f. Fig. 4.7, lanes 7-10, α and β). Extension of the OPTM1-3PPL nascent chain to 204

residues resulted in a loss of any authentic cross-linking to these translocon

components, implying that TM3 has moved out of the translocon by this chain length

(Fig. 4.8B, lanes 7-10). I therefore conclude that the relocation of TM3 from the ER

translocon does not require the presence of the later TM domains (TM4 to TM7) and

appears to be driven solely by chain extension.


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Figure 4. 8 A) A schematic representation of OPTM1-3PPL[cys115] polypeptide chain. Residues 1 to 142 of opsin which contained TM1 to TM3 (in black) were fused to residues 31 to 229 of preprolactin (PPL) (in grey) to generate OPTM1-3PPL. A cysteine probe was located within TM3 at position 115 and is indicated with a white star. B) BMH cross-linking of OPTM1-3PPL[cys115] integration intermediates to translocon components. mRNA encoding OPTM1-3PPL164[cys115] and OPTM1-3PPL204[cys115] was translated in the presence of semi-permeabilised cells and treated as described in the legend of Figure 4.2. Glycosylated and non-glycosylated uncross-linked opsin chains are indicated with (ii) and (i) while truncated nascent chains are marked with asterisks. Authentic adducts to Sec61α and Sec61β are indicated with ‘α’ and ‘β’.


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4.8 Nascent opsin chains are associated with a single copy of the Sec61 complex

during integration

An implicit assumption of site-specific cross-linking analysis exploited above is that the

environment of the nascent chain as a whole is independent of probe location. This

assumption allows a composite model to be generated by combining the results obtained

from the analyses of single cross-linking probes placed at different locations within the

same polypeptide chain. In order to test this assumption, and also to further characterise

the composition of the translocon, opsin nascent chains containing two cysteine probes

were generated. In OP[cys56,115], one cysteine probe is placed at position 56 within

TM1 while the second cysteine probe is at position 115 within TM3 of opsin.

Integration intermediates of OP[cys56,115] of two chain lengths, 164 and 204 residues,

were first synthesised in the presence of semi-permeabilised cells. BMH cross-linking

and immunoprecipitations were then carried out as before (described in section 4.2).

The cross-linking patterns of OP164[cys56,115] and OP204[cys56,115] reflected the

results observed earlier for integration intermediates with only a single cysteine probe

(c.f. Fig. 4.2, lanes 22-25, Fig. 4.7, lanes 7-10, Fig. 4.9, lanes 2-5). Clear adducts were

seen with Sec61α, Sec61β and PAT-10 (Fig. 4.9, lanes 2-5, α, β and P). In addition, a

higher molecular weight adduct of ∼40 kDa was also immunoprecipitated with the α-

Sec61β antibody (Fig. 4.9, lane 5, β+P). This adduct is likely to represent a single

OP164 nascent chain simultaneously cross-linked to both Sec61β (from cys115) and

PAT-10 (from cys56). When the nascent chain is extended to 204 residues, only a single

distinct adduct to PAT-10, most likely from cys56, was observed (c.f. Fig. 4.2, lanes 27-

28, Fig. 4.7, lanes 17-18, Fig. 4.9, lanes 7-8). Therefore, the analysis of nascent chains

with double cysteine probes confirmed the results obtained from the analysis of

integration intermediates with a single cysteine probe, indicating that the nascent chains

as a whole occupy the same environment independent of the cysteine probe location.

In the single cysteine probe analysis of the OP164 integration intermediate, both cys56

and cys115 can individually form adducts with Sec61α (Fig. 4.2, lane 24, Fig. 4.7, lane

9), therefore, if the nascent chain OP164 is adjacent to multiple copies of Sec61α,

having both cys56 and cys115 within a single nascent chain would in principle allow

the integration intermediate to cross-link two copies of the Sec61α subunit at the same


97

time. A glycosylated OP164 nascent chain cross-linked to two molecules of Sec61α is

expected to have a mobility of ∼100 kDa. In fact, no evidence of such a species

containing two copies of the Sec61α subunit cross-linked to a single opsin chain was

observed (Fig. 4.9, lane 4). An adduct of ∼70 kDa was seen, but this adduct contained

both Sec61α and Sec61β, and was also observed in the earlier analysis of

OP164[cys115] (c.f. Fig. 4.9, lanes 4-5, Fig. 4.7, lanes 9-10, (αβ)). These results

indicate that the TM domains of nascent opsin chains are most likely adjacent to only a

single copy of the Sec61α molecule at this stage of biosynthesis. The double probe

studies also showed that PAT-10 is adjacent to the functional Sec61 complex since the

OP164 integration intermediate can be simultaneously cross-linked to both Sec61β and

PAT-10.

Figure 4. 9 BMH cross-linking with OP164 and OP204 integration intermediates containing double cysteine probes. OP164 and OP204 integration intermediates of [cys56,115] were translated in the presence of semi-permeabilised cells and then treated as described in the legend of Figure 4.2. Doubly-glycosylated and non-glycosylated opsin chains which were not cross-linked were indicated with (ii) and (i) respectively. Asterisks denote opsin chains which lack the C-terminal HA tag. Cross-linking products to Sec61α, Sec61β, PAT-10 and Sec61α-Sec61β dimer are indicated with ‘α’, ‘β’, ‘P’ and ‘αβ’ respectively. Nascent chains which were simultaneously cross-linked to Sec61β and PAT-10 were marked with ‘β+P’.


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4.9 Summary

The analysis of the membrane integration of the N-terminal region of opsin revealed

that TM1 interacts with the ER translocon in two discrete phases, each generating a

distinct set of cross-linking partners. In one phase, TM1 is adjacent to both Sec61α and

Sec61β, while in the other phase, TM1 is adjacent to Sec61α and PAT-10. The absence

of TM2 to TM7 of opsin affects TM1 relocation through these phases, delaying the

relocation from the former. The analysis of TM3 integration showed that TM3 engages

the Sec61 complex from a chain length of ∼150 residues and leaves the translocon when

the nascent chain reaches ∼204 residues in length. However, TM3 exit from the

translocon is independent of the presence of TM4 to TM7. Results obtained from the

double probe studies confirmed the observations made with single probes and suggested

that the TM domains of opsin nascent chains associate with only one copy of the Sec61

complex. In addition, analysis of integration intermediates with double cysteine probes

showed that the novel PAT-10 component is in close proximity to a functional Sec61

complex.

Principal conclusions

1) Opsin TM1 experiences at least two distinct phases of Sec61 mediated

environments during opsin biosynthesis. The movement of TM1 through these

phases is dependent on the presence of TM2 and/or TM3.

2) Opsin TM3 exits the translocon as soon as TM4 has been synthesised, but unlike

TM1, relocation of TM3 is independent of the presence of TM4.

3) The TM domains of the nascent opsin chains engage with only a single copy of

the Sec61 complex.

4) PAT-10 is adjacent to a functional Sec61 translocon.

Chapter 5 – The analysis of TM4 to TM7 of opsin

99

CHAPTER 5 Results

The integration of the C-terminal

region of opsin:

TM4 to TM7


100

5.1 Introduction

My examination of the molecular environment of the N-terminal region of opsin,

specifically TM1 to TM3, revealed the distinct behaviour of different TM domains

during their integration at the ER. In this chapter, my study of opsin integration is

further extended to incorporate the remaining opsin TM domains, namely TM4 to TM7.

The prior analysis of TM3 integration had indicated that the ‘gross’ location of a

cysteine probe within the TM domain may influence the pattern of its cross-linking to

translocon components. With this in mind, the molecular environment of at least two

different cysteine probes within each TM domain was initially examined. A unique

cysteine residue was introduced at a specific position within a particular TM domain of

a cysteine-null opsin mutant and a range of integration intermediates of increasing

nascent chain lengths was generated as before (refer to Figure 4.1 in Chapter 4). This

was combined with the use of the sulphydryl-specific homobifunctional cross-linking

reagent BMH to enable a site-specific cross-linking analysis of the proteinaceous

environment of each integration intermediate.

5.2 TM4 exits the translocon upon chain extension

In comparison to some of its other TM domains which are atypically long, opsin TM4 is

a fairly typical transmembrane span, comprising residues 151 to 173 (Palczewski et al.,

2000). Single cysteine probes were placed towards the N-terminus of TM4, at residue

154, and more centrally at residue 165. In the first instance, an integration intermediate

of 204 residues was generated since this was predicted to provide insights into the

environment of TM4 as it is emerging from the ribosome. OP204[cys154] and

OP204[cys165] chains were synthesised in a rabbit reticulocyte translation system in the

presence of digitonin-permeabilised mammalian cells and membrane-integrated

intermediates were isolated by centrifugation. Cross-linking was carried out by

incubation with BMH, with control samples being mock-treated with solvent (DMSO).

The resulting adducts were analysed by immunoprecipitation as previously described.

OP204[cys154] gave a distinct adduct with Sec61β (Fig. 5.1, lanes 2-5, β), but produced

no authentic adducts with Sec61α that were immunoprecipitated with the α-HA

antibody (Fig. 5.1, c.f. lanes 3 and 4, ). A faint ~70 kDa adduct is recognised by both


101

the α-opsin and α-HA antisera (Fig. 5.1, lanes 2 and 3, see X), but this product is not

recovered with the α-Sec61α antiserum (Fig. 5.1, lane 4) and its origin is presently

unknown. OP204[cys165] did not produce discrete adducts with any translocon

components (Fig. 5.1, lanes 7-10). On the basis that OP204[cys154] could be cross-

linked to Sec61β, I conclude the N-terminal region of TM4 has begun to engage the

Sec61 translocon in the 204-residue integration intermediate.

In order to examine the environment of TM4 at different stages during its integration,

BMH-mediated cross-linking was performed with increasing lengths of polypeptide

chains containing the cys154 probe. Samples were treated as before and

immunoprecipitations were carried out with α-opsin, α-HA, α-Sec61α, α-Sec61β and a

non-related (NS) antisera. As observed earlier, cys154 gave an adduct with Sec61β

when the nascent chain is 204 residues (Fig. 5.2, lanes 2-6, β), but upon chain extension

to 259 residues and beyond, no authentic adducts with any translocon components were

observed (Fig. 5.2, lanes 8-12, 14-18, 20-24 and 26-30). Some of the longer chains did

show cross-linking to Sec61β, but none of these species were recognised by the α-HA

antibody and are most likely adducts with shorter nascent chains (Fig. 5.2, c.f. lanes 9

and 12, 15 and 18, 21 and 24, 27 and 30, ). This is particularly apparent where the

cross-linking products are shorter than the nascent chains used for adduct formation

(Fig. 5.2, lanes 24 and 30, see also Meacock et al., 2002).

Products detected with the α-Sec61α antibody were due to the presence of radiolabelled

Sec61α molecules resulting from the translation of endogenous mRNA and these

species do not represent true adducts with Sec61α (Fig. 5.2, •, see Chapter 3.4).

Interestingly, adducts to a ∼10 kDa molecule were observed with the long integration

intermediates, from OP259 to OP357 (Fig. 5.2, lanes 8-9, 14-15, 20-21, 26-27, P?).

Although these adducts were weaker, they resemble the adducts observed between TM1

and PAT-10 (see Chapter 4, Fig. 4.2), implying that TM4 may be adjacent to a PAT-10

like molecule. Taken together, these results imply that TM4 engages the Sec61 complex

at a chain length of 204 residues and moves away from the translocon by the point at

which a chain length of 259 residues has been reached - a stage at which TM5 has been

synthesised. The absence of any adducts with the Sec61 subunits for any of the longer

integration intermediates, OP304 to OP357, indicates that TM4 does not re-associate


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with the translocon during the later stages of opsin biosynthesis (Fig. 5.2, lanes 14-18,

20-24, 26-30).

Figure 5. 1 BMH cross-linking of OP204 integration intermediates from two different single cysteine probes within TM4. OP204[cys154] and OP204[cys165] were synthesised in a rabbit reticulocyte translation system and membrane-integrated products isolated by centrifugation. The samples were treated with BMH (+) or DMSO (-), quenched, and denatured with 1% SDS. Immunoprecipitations using α-opsin, α-HA, α-Sec61α and α-Sec61β antisera were carried out. Uncross-linked doubly-glycosylated and non-glycosylated nascent opsin chains are indicated with (ii) and (i) respectively, while truncated opsin chains which lack the C-terminal HA tag are marked with asterisks. The adduct with Sec61β is indicated with ‘β’. A species containing Sec61α that does not align with the products recognised by the α-HA serum is indicated by ‘ ’ and an adduct with an unknown cross-linking partner by ‘X’.


103

Figure 5. 2 Cross-linking of OP[cys154] integration intermediates of increasing chain lengths to translocon components. Integration intermediates of OP[cys154] were synthesised by in vitro translation and subjected to BMH cross-linking and immunoprecipitation as described in the legend to Figure 5.1. Endogenous, radiolabelled Sec61α molecules are indicated with ‘•’, while Sec61β-containing adducts to truncated opsin chains lacking the C-terminal HA tag are marked with ‘ ’. Putative adducts to a PAT-10 like molecule are labelled ‘P?’. Other symbols are as previously defined in the legend to Figure 5.1.


104

5.3 Opsin TM5 engages the translocon at 259 residues

In order to examine the integration of opsin TM5 (residues 200 to 225, Palczewski et

al., 2000) into the lipid bilayer by a similar cross-linking approach, single cysteine

probes were introduced at three different locations within TM5, namely residue 204

near its N-terminus, residue 217 near its centre, and residue 229 at the C-terminal

boundary of TM5. Integration intermediates of 259 residues, a length at which TM5 is

predicted to be largely out of the ribosome, were generated and their proximity to

components of the ER translocon examined by cross-linking as previously described.

Interestingly, in this case, only the nascent chain with a cysteine probe at the boundary

of TM5, OP259[cys229], generated authentic cross-linking adducts with Sec61

components, i.e. the Sec61β subunit (Fig. 5.3, see lanes 12-15, β). Whilst there may also

be some cross-linking to Sec61α from the OP259[cys229], this was certainly not

unambiguous and there was no compelling evidence of any such product in the α-HA

recovered material (Fig. 5.2, lane 13, ∼66 kDa range). In contrast, the OP259[cys204]

and OP259[cys217] showed no authentic adducts with subunits of the Sec61 complex

(Fig. 5.3, lanes 3-5 and 8-10), and only endogenously encoded Sec61α (Fig. 5.3, lanes 4

and 9, •) and incomplete chains cross-linked to Sec61β (Fig. 5.3, lane 10, ) were seen.

Since cys229 is adjacent to the Sec61β subunit in the 259 residue integration

intermediate, I conclude that opsin TM5 has engaged the Sec61 translocon at this stage

of biosynthesis.


105

Figure 5. 3 BMH cross-linking of OP259 integration intermediates using different TM5 specific probes. mRNA chains encoding OP259[cys204], OP259[cys217] and OP259[229] were translated in vitro in the presence of semi-permeabilised cells. BMH cross-linking and immunoprecipitations were performed as described in the legend to Figure 5.1. Uncross-linked doubly-glycosylated and non-glycosylated opsin chains are indicated with (ii) and (i) respectively. Truncated opsin chains which lack the C-terminal HA tag are marked with asterisks. Endogenous radiolabelled Sec61α molecules are denoted with ‘•’ and a Sec61β-containing adduct to truncated nascent opsin chains is marked with ‘ ’. Adducts with Sec61β are indicated with ‘β’.


106

5.4 Opsin TM5 is adjacent to a PAT-10-like molecule during its integration

The analysis of the C-terminal region of opsin requires long integration intermediates to

be generated for cross-linking, since the bulk of the polypeptide must be synthesised to

expose this part of the protein. As is readily apparent from the analysis of TM5 shown

above, adducts obtained with long nascent chains are often very weak and diffuse (Fig.

5.3, see lanes 3, 8 and 13). In addition, significantly more products lacking the C-

terminal HA tag were present (Fig. 5.3, lanes 2, 7 and 12, *, see also Chapter 3.2). As a

consequence, the interpretation of the resulting cross-linking products can be difficult.

In order to facilitate the analysis of TM5 to TM7 of opsin, a truncated version of the

opsin polypeptide chain was generated in which the first 35 residues of opsin, which

contains the two N-glycosylation sites, were fused to a C-terminal region of opsin

containing TM5 to TM7 (residues 195 to 348) to form the OPN/5-7 polypeptide chain

(Fig. 5.4A). In vivo expression has shown that opsin TM5 to TM7 (residues 195 to 348)

is one of several fragments that behaves as an ‘independent folding domain’ (Ridge et

al., 1996). The potential advantage of this deletion mutant is that shorter integration

intermediates can be used for the cross-linking analysis of TM5 to TM7.

In the first instance, the topology of the OPN/5-7 polypeptide chains was ascertained by

determining if cross-linking products were formed with doubly glycosylated nascent

chains. An integration intermediate of OPN/5-7 with a cysteine probe in the first TM

domain (OPN/5-7 259[cys229]) was subjected to BMH cross-linking and

immunoprecipitation, followed by treatment with endoglycosidase H to remove the

asparagine-linked glycan groups. BMH-dependent adducts to Sec61α and a ∼20 kDa

unidentified molecule were observed (Fig. 5.4C, lanes 2-4, α and X) and these adducts

were endoglycosidase H sensitive, as shown by the faster migration of the products

(Fig. 5.4C, lanes 7-9, α and X), implying that these adducts were formed with OPN/5-7

nascent chains which were glycosylated. This indicated that OPN/5-7 polypeptide

chains have been inserted in the correct orientation into the ER membrane.


107

Figure 5. 4 (A) A schematic diagram of the OPN/5-7 polypeptide chain. The first 35 residues of opsin was fused to a C-terminal region of opsin (residues 195 to 348) which contains TM5, TM6 and TM7 (TM domains are shown in black). (B) Predicted topology of OPN/5-7 polypeptide chain. TM domains are shown as grey boxes while the numbers indicate the position of TM5 to TM7 in context of the OPN/5-7 polypeptide chain. N-glycosylation sites are marked with ‘Y’ and the C-terminal HA tag is represented with a grey line. (C) Cross-linking adducts are formed with glycosylated OPN/5-7 nascent chains. An integration intermediate of OPN/5-7 with a single cysteine probe in TM5 was subjected to BMH mediated cross-linking and immunoprecipitation as described in legend 5.1. Duplicate samples were additionally treated with endoglycosidase H to cleave the asparagine-linked glycan groups (+ Endo H). Adducts with Sec61α are labelled ‘α’ and other distinct BMH dependent cross-linking adducts are marked with ‘X’. Other symbols are as previously defined in Figure 5.1.


108

However, whilst the topology of TM5 to TM7 should be the same as in the wild type

protein (Fig. 5.4B), the relative position of TM5, TM6 and TM7 in the context of the

polypeptide chain has changed (i.e. TM5 is now the first TM domain and so on). For

this reason, a cross-linking analysis with the normal opsin polypeptide chain was

performed concurrently with that of OPN/5-7 to compare the two species. Although the

analysis of TM5 integration should ideally utilise a cysteine probe located within the

TM domain, earlier analysis of cys204 and cys217 introduced into TM5 failed to

produce any adducts even at the shortest chain length (OP259) (see section 5.3). Hence,

further analysis of TM5 was carried out using integration intermediates with a cysteine

residue at position 229. Cys229 is located in the hydrophilic loop just at the C-terminal

boundary of TM5, thus, cross-linking performed with integration intermediates of the

cys229 mutant should reflect the environment of at least the C-terminal end of the TM

domain.

Four integration intermediates of full length opsin, OP259[cys229], OP304[cys229],

OP339[cys229] and OP357[cys229], were generated by in vitro translation and

subjected to BMH-mediated cross-linking and immunoprecipitation as before. As

described earlier in section 5.3, OP259[cys229] formed adducts with Sec61β (Fig. 5.5A,

lanes 2-6, β). In this experiment, it also appeared that there may be an authentic adduct

with Sec61α (Fig. 5.5A, c.f. lanes 3 and 5, α). At the longer chain lengths of 304, 339

and 357 residues, Sec61α- and Sec61β-containing adducts were present to some degree

but it was impossible to determine whether these adducts were immunoprecipitated with

the α-HA antibody (Fig. 5.5A, c.f. lanes 9 and 11, 15 and 17, 21 and 23, α? and β?).

What was clear was that distinct adducts with a ∼10 kDa protein were observed with

each of these longer nascent chains (Fig. 5.5A, lanes 8-9, 14-15, 20-21, P). This adduct

is remarkably similar to the species observed between a TM1 cysteine probe and PAT-

10 (see Chapter 4.2, Fig. 4.2), implying that TM5 is adjacent to a PAT-10-like

molecule.

When a similar cross-linking analysis was performed with the equivalent OPN/5-7

intermediates, adducts with Sec61 subunits were more distinct. Hence, the analysis of

OPN/5-7 259[cys229] revealed a prominent adduct to Sec61α, which was clearly

recognised by the α-HA antibody (c.f. Fig. 5.5A, lanes 3 and 5 to Fig. 5.5B, lanes 3 and

4, α). Similarly, when longer OPN/5-7 integration intermediates were examined, distinct


109

adducts to Sec61α were observed even until the longest chain length (Fig. 5.5B, lanes 7-

9, 12-14, 17-19, α). Cross-linking to a ∼10 kDa protein from cys229 was also seen from

OPN/5-7 304 until OPN/5-7 357 (Fig. 5.5B, lanes 7-8, 12-13, 17-18, P).

Figure 5. 5 BMH cross-linking of cys229 integration intermediates of A) normal-length opsin polypeptide chain, and B) OPN/5-7 polypeptide chain. OP[cys229] nascent chains of 259, 304, 339 and 357, and OPN/5-7 integration intermediates of equivalent lengths, were synthesised by in vitro translation and subjected to BMH cross-linking and immunoprecipitations as described in the legend to Figure 5.1. Putative adducts to Sec61α and Sec61β are denoted with ‘α?’ and ‘β?’ respectively, while clear adducts to Sec61α and Sec61β are indicated with ‘α’ and ‘β’. PAT-10 adducts are marked with ‘P’ and putative adducts with ribosomal proteins are labelled ‘R’ (see Laird and High, 1997). Other symbols are as defined in the legend to Figure 5.3.


110

OPN/5-7 259[cys229] gave a discrete adduct to Sec61β (Fig. 5.5B, lanes 2-5, β).

However, for the longer OPN/5-7[cys229] nascent chains, it is difficult to ascertain

whether the faint Sec61β adducts that were still observed, were formed with authentic

integration intermediates, since the PAT-10 adducts mask the presence of any Sec61β

adducts due to their similar molecular weights (Fig. 5.5B, lanes 10, 15, 20, β?). On the

basis of adduct formation with Sec61α, I concluded that TM5 engages the Sec61

complex at 259 residues and remains associated with the translocon until the entire

opsin polypeptide chain has been synthesised. TM5 is also adjacent to a PAT-10-like

molecule from 304 residues and remains in close proximity to it throughout the

remainder of nascent chain synthesis.

5.5 Opsin TM1 and TM5 are adjacent to a single copy of PAT-10

One of the striking features of the behaviour of opsin TM1 is that it associates with the

novel ∼10 kDa protein, PAT-10, as soon as the nascent chain is long enough to allow

the next TM domain (TM2) to emerge from the ribosome, and this interaction lasts for a

prolonged period of opsin synthesis (see Fig. 4.2) and probably until the completion of

opsin biosynthesis and chain termination (Meacock et al., 2002). Surprisingly, TM5 is

also adjacent to a ∼10 kDa protein at a stage when the subsequent TM domain (TM6)

has been synthesised, and adduct formation is observed for all the longer chains

analysed as integration intermediates. As the identity of PAT-10 is unknown, it is

difficult to ascertain if TM5 is interacting with PAT-10 or a distinct ∼10 kDa molecule

with similar properties. In order to better establish whether TM5 is adjacent to PAT-10,

and to determine whether the nascent opsin chain associates with multiple copies of

identical or different ∼10 kDa proteins, an OP304 integration intermediate with one

cysteine probe in TM1 (cys56) and a second cysteine probe flanking TM5 (cys229) was

generated. A chain length of 304 residues was chosen because it is the shortest

integration intermediate which would allow both TM1 and TM5 to cross-link the ∼10

kDa protein (see Fig. 5.5A, P).

BMH-mediated cross-linking of OP304[cys56,229] produced only one major adduct

which corresponded to the nascent chain being cross-linked to a single copy of a ∼10

kDa protein (Fig. 5.6, lanes 2-3, P). If the OP304 integration intermediate were to cross-

link two molecules of the ∼10 kDa protein from both TM1 and TM5, then a higher


111

molecular weight adduct of ∼50 kDa would be observed. Since only one copy of the

∼10 kDa protein is adjacent to OP304[cys56,229], the protein found adjacent to TM5 is

presumably the same molecule that is adjacent to TM1. Hence, I conclude that TM5 is

most likely associated with PAT-10, and that there is only one copy of PAT-10 adjacent

to the functional Sec61 translocon.

Figure 5. 6 Double probe analysis of the OP304 integration intermediate. mRNA transcripts encoding OP304[cys56,229] nascent chains were translated in vitro in the presence of semi-permeabilised cells. Membrane-integrated products were subjected to BMH cross-linking and immunoprecipitations as described in the legend to Figure 5.1. Uncross-linked doubly-glycosylated and non-glycosylated nascent opsin chains are labelled (ii) and (i), while truncated nascent chains are marked with an asterisk. The presumptive adduct to PAT-10 is indicated with ‘P’.


112

5.6 Opsin TM6 engages the translocon throughout opsin biosynthesis

The molecular environment of TM6 during opsin biosynthesis was initially examined

by introducing a unique cysteine probe at two separate locations within the TM domain,

namely residue 254 in the middle of TM6 or residue 275 near the C-terminal end of the

TM domain. For both cysteine positions, integration intermediates of 304 residues in

which TM6 is presumed to have fully emerged from the ribosome, were generated by in

vitro translation. BMH-mediated cross-linking and immunoprecipitations were carried

out as previously described in section 5.2.

Whilst OP304[cys275] gave a distinct adduct with Sec61β that was also recognised by

the α-HA antibody (Fig. 5.7, lanes 7-10, β), it was not clear that the Sec61β adduct seen

with OP304[cys254] was recognised by the α-HA antibody and hence uncertain as to

whether it was formed with the 304 residue long chain in this case (Fig. 5.7, lanes 2-4,

β?). Likewise, whilst products were immunoprecipitated with the α-Sec61α serum in

both cases (Fig. 5.7, lanes 4 and 9, •), no obvious equivalent was seen in the

accompanying α-HA immunoprecipitation (Fig. 5.7, lanes 3 and 8). Thus, with such

long intermediates, only evidence of proximity between the C-terminal region of TM6

and the Sec61β subunit could be unequivocally determined.

Since cys275 gave clear and unambiguous data, integration intermediates of three

different lengths, 304, 339 and 357, each with a single cysteine at residue 275, were

analysed by BMH-mediated cross-linking. As before, OP304[cys275] gave a distinct

adduct with Sec61β, indicating that TM6 has engaged the Sec61 complex (Fig. 5.8A,

lane 2-6, β). Extending the nascent chain to 339 residues allowed TM6 to form discrete

adducts with both Sec61α and Sec61β (Fig. 5.8A, lanes 8-12, α and β). Upon further

extension to OP357, an authentic adduct with Sec61α appeared to be maintained (Fig.

5.8A, lanes 14, 15 and 17, α) whilst the authenticity of a potential Sec61β adduct was

less obvious (Fig. 5.8A, lanes 14, 15 and 18). As seen for the analysis of TM5 (section

5.4), adducts with long nascent chain lengths were diffuse and difficult to substantiate

by parallel immunoprecipitation using the α-HA sera. In order to try to resolve some of

the ambiguities resulting from the cross-linking of these longer chains, a parallel

analysis was performed using the opsin deletion mutant, OPN/5-7. This analysis

revealed a similar, albeit more clear cut, pattern of adduct formation with Sec61


113

subunits. Thus, an adduct with Sec61β was seen for the shortest integration

intermediate, OPN/5-7 304[cys275], while adducts with both Sec61α and Sec61β were

observed with OPN/5-7 339[cys275] and OPN/5-7 357[cys275] (Fig. 5.8B, lanes 2-5, 7-

10, 12-15, α and β). Taken together, these results suggest that TM6 engages the Sec61

complex when the nascent chain is 304 residues long and remains associated with the

translocon throughout the remainder of opsin synthesis. Since the cross-linking pattern

to translocon components changes at increasing chain lengths, it is plausible that TM6

relocates from its initial location within the Sec61 complex during the extension of the

nascent chain, along the lines of the process previously described for TM1.

Figure 5. 7 BMH dependent cross-linking from single cysteine probes in opsin TM6. In vitro translation of OP304[cys254] and OP304[cys275], BMH cross-linking and immunoprecipitation were carried out as described in the legend of Figure 5.1. Uncross-linked doubly-glycosylated, non-glycosylated and truncated opsin chains are indicated with (ii), (i) and ‘ ’ respectively. Distinct and putative adducts to Sec61β are labelled ‘β’ and ‘β?’ respectively. Presumptive endogenous radiolabelled Sec61α molecules are marked ‘•’.


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Figure 5. 8 BMH cross-linking of cys275 integration intermediates of A) normal-length opsin polypeptide chains, and B) OPN/5-7 polypeptide chains. OP304[cys275], OP339[cys275], OP357[cys275] and their respective OPN/5-7 equivalents, were synthesised by in vitro translation in the presence of semi-intact cells and subjected to BMH cross-linking and immunoprecipitations as described in the legend of Figure 5.1. Adducts to Sec61α and Sec61β are labelled ‘α’ and ‘β’, while nascent chains simultaneously cross-linked to both Sec61α and Sec61β are indicated with ‘αβ’. Unidentified adducts are marked with ‘X’. Other symbols used were previously defined in the legend of Figure 5.3.


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5.7 Opsin TM7 is associated with the translocon throughout opsin biosynthesis

In the analysis of the final TM domain, opsin TM7, the cross-linking patterns of

cysteine probes at two different positions were again analysed. One cysteine probe was

located near the N-terminus of TM7 (residue 287) and the other near its C-terminus

(residue 308). Integration intermediates with a nascent chain length of 339 residues,

OP339[cys287] and OP339[cys308], were then generated and subjected to BMH cross-

linking and immunoprecipitation.

Both OP339[cys287] and OP339[cys308] gave only distinct adducts with Sec61β that

also appeared to be recognised by the α-HA serum (Fig. 5.9, lanes 3 and 5, 8 and 10, see

β). The behaviour of the two TM7 probes with respect to adduct formation with Sec61α

was more difficult to assess (Fig. 5.9, lanes 3 and 4, 8 and 9). Probe 287 was selected

for further analysis, and initially two different intermediates investigated, namely

OP339[cys287] and OP357[cys287]. In both cases, adducts with Sec61β were observed

and appeared to be authentic by the criteria of α-HA immunoprecipitation (Fig. 5.10A,

lanes 3 and 5, 8 and 10, β). As before, the presence of adducts with Sec61α could not be

unambiguously distinguished (Fig. 5.10A, lanes 3 and 4, 8 and 9, α?). In order to better

understand TM7 integration, equivalent integration intermediates were analysed for the

OPN/5-7 polypeptide. As with the normal length chains, authentic adducts with Sec61β

were seen (Fig. 5.10B, lanes 3 and 5, 8 and 10, β). However, in this case, adducts with

Sec61α could also be definitely identified (Fig. 5.10B, lanes 3 and 4, 8 and 9, α). Taken

together, these data indicate TM7 engages the Sec61 translocon after emerging from the

ribosome and, under normal circumstances, remains associated with the ER translocon

for the remainder of opsin biogenesis.


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Figure 5. 9 BMH mediated cross-linking from distinct cysteine probes in opsin TM7. In vitro translation of OP339[cys287] and OP339[cys308], BMH cross-linking and immunoprecipitations were carried out as described in the legend of Figure 5.1. Symbols indicated were as previously defined in the legend of Figure 5.3.


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Figure 5. 10 BMH cross-linking of cys287 integration intermediates of A) normal-length opsin polypeptide chain, and B) OPN/5-7 polypeptide chain. OP339[cys287], OP357[cys287] and their OPN/5-7 equivalents, were synthesised using a rabbit reticulocyte translation system in the presence of semi-permeabilised cells and treated with BMH as described in the legend of Figure 5.1. Symbols used were defined in the legend of Figure 5.8.


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5.8 Summary

The C-terminal region of opsin, comprising TM4 to TM7, experiences distinct

molecular environments during their integration into the ER membrane. TM4 engages

the Sec61 complex as TM3 departs and is itself then ‘displaced’ as TM5 emerges from

the ribosome. In contrast, TM5, TM6 and TM7 all appear to remain in close proximity

to the ER translocon until the entire opsin polypeptide chain has been synthesised.

Interestingly, like TM1, TM5 is found adjacent to a component assumed to be PAT-10

during opsin biosynthesis. A double probe analysis indicated that only one copy of

PAT-10 is in close proximity to an integrating opsin polypeptide chain. The analysis of

the C-terminal TM domains of opsin was facilitated by the use of a shorter version of

opsin, OPN/5-7, which allowed unambiguous interpretation of the cross-linking data.

Conclusion

1) Opsin TM4 enters the Sec61 complex as TM3 departs, and then exits the

translocon itself once TM5 emerges from the ribosome.

2) Opsin TM5, TM6 and TM7 remain associated with the translocon throughout

polypeptide chain synthesis.

3) Both TM1 and TM5 of a single opsin chain are adjacent to a single copy of

PAT-10, suggesting that only one PAT-10 molecule is associated with a

functional ER translocon.

Chapter 6 – Probing the environment of a translocating opsin chain

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CHAPTER 6 Results

Probing the environment of a

translocating nascent opsin chain


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6.1 Introduction

The previous chapters have focused on the identity of the ER translocon components

that are adjacent to specific TM domains of opsin during their integration into the

membrane. The distinct behaviour of each TM domain was demonstrated by their

different cross-linking patterns to subunits of the Sec61 complex and other ER

components, particularly PAT-10. One limitation of this approach was that the

relationship of TM domains to the lipid bilayer was not directly analysed. Hence, loss of

cross-linking to Sec61 subunits was taken to reflect a transfer to the phospholipid phase

in line with several previous studies (Martoglio et al., 1995; McCormick et al., 2003;

Mothes et al., 1997). In order to obtain an alternative view of TM environment during

opsin polypeptide chain elongation, a distinct approach to analyse the location of single

cysteine probes in opsin integration intermediates was explored.

Figure 6. 1 Structures of (a) 4-acetamido-4′-maleimidylstilbene-2-2′-disulfonic acid (AMS) and (b) QSY 9 C5-maleimide (QSY). Both AMS and QSY are sulphydryl specific modification reagents. Their molecular weight (MW) and their solubility properties are indicated.

AMS MW: 536.44

Water solubility: High

Lipid solubility: Low

QSY MW: 1083.3

Water solubility: Moderate

Lipid solubility: High

(a)

(b)


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In this chapter, two monofunctional sulphydryl-specific reagents were used to examine

the ‘environment’ of TM3 during its movement from the ribosome into the ER

translocon. 4-acetamido-4′-maleimidylstilbene-2-2′-disulfonic acid (AMS) is a

hydrophilic molecule (Fig. 6.1), and thus reacts preferentially with free sulphydryl

groups in a hydrophilic environment, such as the cytosol, the aqueous translocon pore or

the lumen of the ER (Krishnasastry et al., 1994). On the other hand, QSY 9 C5-

maleimide (QSY) is a more hydrophobic molecule (Fig. 6.1) that would be predicted to

react with free sulphydryl groups in both hydrophilic and hydrophobic environments

including the phospholipid bilayer of the ER membrane. The rationale for this work was

to explore whether there was any correlation between the accessibility of a particular

cysteine residue to these two probes and the stage of opsin biogenesis that was explored.

As with the site-specific cross-linking analysis, this ‘probe accessibility’ study took

advantage of single cysteine residues located within the opsin chain. In this instance, I

focused on analysing opsin TM3 which I had previously found to exit the ER translocon

independently of TM4 to TM7 (see Chapter 4). Thus, integration intermediates of

increasing chain lengths were generated to mimic different stages of TM3 insertion and

the access of AMS and QSY to different cysteine probes present in these nascent chains

was determined.

6.2 AMS and QSY can modify cysteine probes in the ER lumen

Previous studies have shown that small molecules of ∼0.5 kDa can readily cross the ER

membrane and access the lumen (Le Gall et al., 2004). The movement of these

molecules is passive, although their mode of entry is not known. Since AMS and QSY

have molecular weights of ∼0.5 kDa and ∼1 kDa respectively, the permeability of the

ER membrane to AMS and QSY was first determined. An opsin integration

intermediate of 96 residues with a single cysteine probe at position 14 near its N-

terminus was generated for this purpose. At 96 residues, the nascent chain is long

enough to allow the full translocation of the opsin N-terminus into the ER lumen and it

is therefore N-glycosylated at both available sites (c.f. Fig. 4.2). Hence, cys14 can only

be modified by AMS and QSY if these reagents traverse the ER membrane.

Modification of the cysteine probe with AMS or QSY should result in a small increase

in molecular weight of the nascent chain, producing an alteration in the migration of the

product up on SDS PAGE.


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mRNA transcripts encoding OP96[cys14] were translated in vitro in a rabbit

reticulocyte translation system supplemented with digitonin-permeabilised mammalian

cells, and the membrane fraction was isolated by centrifugation. The membrane-

integrated products were incubated with either AMS or QSY, or mock treated with

solvent (DMSO) only. The reaction was quenched by the addition of β-mercaptoethanol

and the samples were denatured with 1% SDS. The products were then recovered by

immunoprecipitation with the α-HA antiserum which recognised the C-terminal epitope

tag present in all the intermediates studied. Since modification with AMS or QSY was

expected to produce only small shifts in mobility on SDS PAGE, duplicate samples

were treated with endoglycosidase H to cleave the glycan groups attached to correctly

integrated opsin chains and increase any alteration in apparent mobility due to

modification.

This preliminary experiment showed that a shift in mobility could be observed for both

the AMS- and QSY-treated samples (Fig. 6.2a and b, c.f. lanes 1 and 2, 3 and 4). Most

importantly, these shifts were observed for the doubly-glycosylated opsin chains,

indicating that properly integrated nascent chains have been modified by AMS and

QSY (Fig. 6.2a, lanes 2 and 4, (ii)). Treatment of the nascent chains with

endoglycosidase H produced a more apparent change in the mobility of modified

polypeptide chains (Fig. 6.2b, c.f. lanes 1 and 2, 3 and 4). A comparison of the opsin

chains after endoglycosidase H treatment also suggested that the majority of the

polypeptides were modified by AMS and QSY, indicating that the environment of the

bulk of the nascent chains would be reported by this approach. Since cys14 is present in

the ER lumen, these results indicate that both AMS and QSY can cross the ER

membrane and react with sulphydryl groups present in the lumen. This behaviour

should allow ready access of both AMS and QSY to cysteine residues present in regions

of the polypeptide that are located in an aqueous environment in the context of an

integration intermediate (c.f. cartoon to the side of Figure 6.2).


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Figure 6. 2 AMS and QSY modification of OP96[cys14] integration intermediates. A schematic representation of the ribosome-nascent chain complex and the translocon is shown in the right panel. The translocon is represented by dark grey boxes, the TM domain is represented by a black box and the position of the cysteine residue is indicated by a star. The two glycosylation sites on the opsin chain are marked with ‘Y’. Membrane-associated OP96[cys14] integration intermediates were isolated as previously described (Chapters 4 and 5) and treated with AMS (+) or QSY (+), or mock-treated with DMSO (-). The reactions were quenched with β-mercaptoethanol, the samples were denatured with 1% SDS, and reaction products recovered by immunoprecipitation with the α-HA antiserum. Panel A shows samples as recovered, panel B shows samples after endoglycosidase H treatment to remove N-linked glycans. Doubly-glycosylated and non-glycosylated opsin chains are indicated with (ii) and (i) respectively.


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6.3 Opsin nascent chains are modified by AMS in the absence of membranes

Having established proof of principle and shown that both AMS and QSY have ready

access to a cysteine probe in the lumenal side of the ER membrane, I next established

whether these reagents were suitable for the analysis of cysteine probes located in TM3.

To this end, single cysteine probes were introduced at three different locations relative

to TM3. The first probe was located at residue 107 within the hydrophilic loop

preceding TM3. A second unique cysteine was introduced at residue 115 to allow the

environment of the N-terminal region of TM3 to be monitored, while a third cysteine

residue was introduced at residue 124 to report the middle of TM3.

In the first instance, I established whether these different cysteine probes could be

modified in the absence of ER membranes such that a visible shift in mobility upon

SDS PAGE could be seen. Thus, polypeptide chains of four different lengths, OP130,

OP140, OP150 and OP164 were examined for each cysteine mutant. The exception is

cys124, where an OP130 chain could not be generated because the cysteine probe is so

close to the C-terminus that it is replaced by the 9 residue HA tag. The position of TM3

and the cysteine probes relative to the ribosome for each nascent chain length is

represented schematically in Figure 6.3. At the shortest chain length examined (130

residues), TM3 is only partly synthesised and the cysteine probes are predicted to be

still buried in the ribosome (Kowarik et al., 2002). As the nascent chains become

progressively longer, TM3 is expected to move out from the ribosome and by 164

residues, TM3 should have largely emerged. The generation of polypeptide chains with

these precise lengths allows the ‘environment’ of the single cysteine probes to be

monitored during nascent chain elongation. As a further control, cysteine-null chains for

each of the intermediates were also generated and subjected to identical treatments.


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Figure 6. 3 AMS modification of various OP130 to OP164 chains. In the schematic diagrams (top and right), TM3 is represented by a black box while the numbers indicate the relative positions of the single cysteine probes (star). Regions of the nascent chain which are not shown in the diagram are indicated with dotted lines. Nascent chains of 130 (a), 140 (b), 150 (c) or 164 residues (d) with single cysteine probes as indicated, were synthesised in vitro in the absence of membranes. Ribosome-nascent chain complexes were isolated and treated with either AMS (+) or DMSO (-). The reaction was quenched with β-mercaptoethanol and products recovered by immunoprecipitation using α-HA antiserum.


126

Cysteine-null polypeptide chains that contain no thiol groups showed no changes in

apparent molecular mass after incubation with AMS, indicating that the AMS reaction

is cysteine-specific (Fig. 6.3 panels a to d, lanes 7 and 8). When the nascent chain is 130

residues long, a small proportion of the OP130[cys115] intermediates showed a reduced

mobility (Fig. 6.3a, lanes 3 and 4). This implies that AMS can enter the ribosome tunnel

since cys115 is expected to be within the ribosome at 130 residues. However, in this

location, the majority of cysteine residues were not affected. A larger proportion of

OP130 chains were modified when the cysteine probe is placed at position 107,

implying greater AMS accessibility to residues nearer to the exit site of the ribosomal

tunnel (Fig. 6.3a, lanes 5 and 6). When longer polypeptide chains, OP140, OP150 and

OP164, were analysed, all the nascent chains exhibit a slower mobility in the presence

of AMS, irrespective of the location of the cysteine probe (Fig. 6.3b, c, d, lanes 1-6).

This shows that the single cysteine probes in the nascent chains can be modified by

AMS when present in a hydrophilic environment located near to or beyond the

ribosomal exit site.

6.4 The environment of cys124 in the OP150 and OP164 integration intermediates

is altered in the presence of the ER translocon

During the integration of opsin TM3 into the ER membrane, this region of polypeptide

must leave the ribosomal exit tunnel, enter the ER translocon, and finally relocate into

the lipid bilayer. In order to better understand this process, the access of AMS to TM3

was now re-investigated in the context of ribosome bound membrane integration

intermediates identical to those previously used for cross-linking studies. Different

stages of biosynthesis were represented by integration intermediates of increasing

lengths and the location of the cysteine probes was analysed as previously established

(c.f. Fig. 6.3). Experimental details were as previously described except that nascent

chains were synthesised in the presence of ER derived membranes allowing N-

glycosylation. For this reason, duplicate samples were treated with endoglycosidase H

to remove the glycans and enhance small mobility shifts (c.f. Fig. 6.2).

No shifts in mobility were observed with any of the cysteine-null nascent chains

confirming the specificity of AMS modification (Fig. 6.4a, b, c and d, lanes 7 and 8). As

before (section 6.3), only a small proportion of OP130 chains with a cys115 probe gave


127

a molecular weight shift consistent with this residue being deep within the ribosome

(Fig. 6.4a, lanes 3 and 4). A much larger proportion of OP130 chains with a cys107

probe were modified, consistent with greater AMS accessibility as the probe is located

closer to the exit site (Fig. 6.4a, lanes 5 and 6).

At 140 residues, cysteine probes in all locations gave a shift in molecular weight due to

AMS modification (Fig. 6.4b, lanes 1-6). This was especially obvious after the

endoglycosidase H treatment and indicates that all of the cysteines are in a hydrophilic

environment. At this stage, cys107 may have emerged from the ribosome and have

entered the aqueous environment of the translocon pore, while cys115 and cys124 are

most likely still within the ribosomal tunnel. Clearly, by whatever route, AMS has ready

access to all three cysteine probes in the OP140 integration intermediate. Strikingly,

when the nascent chains were extended to 150 and 164 residues, only polypeptide

chains with cys107 and cys115 probes now produced an AMS-mediated mobility shift,

while nascent chains with cys124 displayed no change in apparent molecular weight

(Fig. 6.4c and d, c.f. lanes 1 and 2, to lanes 3-6). The simplest interpretation of this

result is that at these chain lengths, cys107 and cys115 have remained in an aqueous

environment, while cys124 has relocated to an environment that is not accessible to

AMS (c.f. cartoons to the right of Figure 6.4).


128

Figure 6. 4 AMS modification of various OP130 to OP164 integration intermediates. The integration intermediates were synthesised and treated with AMS as described in the legend to Figure 6.3. Duplicate samples were treated with endoglycosidase H (+ EndoH) to remove N-linked glycans.


129

6.5 Cys124 in OP150 and OP164 integration intermediate is in a hydrophobic

environment

The absence of AMS modification for OP150[cys124] and OP164[cys124] suggests that

cys124 may be in a location that cannot be modified by AMS, for example, a

hydrophobic environment. Alternatively, the extension of the nascent chain from 140 to

150 or 164 residues may alter the nascent chain to fold into a conformation that

prohibits any modification of cys124. In order to distinguish between such possibilities,

the integration intermediates were analysed with a second sulphydryl specific reagent,

QSY. Since QSY is moderately hydrophobic, it should react with cysteine residues in

both hydrophobic and hydrophilic environments. If QSY can react with cys124 in

OP150 and OP164, this would suggest that it is the environment of the cysteine probe

that prevents its AMS modification and not the conformation of the nascent chain.

Identical integration intermediates to those shown in Figure 6.4 were treated with QSY

and the products analysed as before. As for AMS, the cysteine-null integration

intermediates were not modified with QSY, showing that QSY reaction is cysteine

specific (Fig. 6.5a, b, c and d, lanes 7 and 8). For OP130, a molecular weight shift was

observed for both cys107 and cys115, indicating that QSY can enter the ribosome (Fig.

6.5a, lanes 3-6). QSY modifications were also seen for all three cysteine residues in the

OP140 integration intermediate as indicated by changes in mobility similar to those

previously observed with AMS (Fig. 6.5b, lanes 1-6, see also Fig. 6.4b, lanes 1-6). As

previously observed, the shifts were more apparent after endoglycosidase H treatment

(Fig. 6.5b, +Endo H, lanes 1-6).


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Figure 6. 5 QSY modification of various OP130 to OP164 integration intermediates. The integration intermediates were synthesised and treated with QSY as described in the legend to Figure 6.4.


131

When the OP150 and OP164 intermediates were analysed, cys107 and cys115

continued to be modified by treatment with QSY (Fig. 6.5c and d, lanes 3-6). Most

significantly however, QSY was found to react with cys124 in these longer integration

intermediates (Fig. 6.5c and d, lanes 1 and 2). This indicates that at nascent chain

lengths of 150 and 164 residues, cys124 is most likely in a hydrophobic environment

which permits QSY access to the cysteine probe but excludes the hydrophilic AMS.

Since cys124 is in the middle of TM3, it is possible that this region of the nascent chain

either contacts phospholipids or is embedded in a hydrophobic region of the Sec61

complex (refer to the schematic diagrams of the ribosome-nascent chain complex at the

translocon in Fig. 6.5c and d).

6.6 The loop region C-terminal to TM3 is in a hydrophilic environment at 164

residues

An alternative explanation of the inaccessibility of cys124 to AMS in the OP150 and

OP164 intermediates is that some type of ‘gating’ event occurs at these chain lengths

(Alder & Johnson, 2004) and prevents access of AMS via closure of the Sec61

translocon and/or the tight binding of the ribosome to the Sec61 complex. Such gating

would also prevent access to probes C-terminal of cys124 that were in a hydrophilic

environment, and this possibility was investigated further to better establish the basis for

the specific absence of AMS modification. To this end, a unique cysteine was

introduced at residue 140 of opsin to determine the environment of the hydrophilic

region of polypeptide C-terminal of TM3. Since in OP164, cys140 is predicted to be

near the ribosomal exit site, an additional integration intermediate of 174 residues was

also generated to examine the environment of cys140 when it was predicted to have

emerged from the ribosome.

In the first instance, OP164[cys140] and OP174[cys140] were synthesised in the

absence of membranes to ascertain that cys140 can be modified by AMS and QSY

when in a freely accessible aqueous environment. Both OP164[cys140] and

OP174[cys140] displayed shifts in molecular weight after treatment with AMS or QSY,

indicating that cys140 can be modified by both reagents at both chain lengths (Fig. 6.6a

and b, lanes 1 and 2, 3 and 4). Having established that cys140 could be modified in the


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context of OP164 and OP174 chains, the analysis was repeated using membrane

integration intermediates where the nascent chains were trapped in the ER translocon.

Figure 6. 6 AMS and QSY modification of cys140 in (a) OP164 and (b) OP174 chains. mRNA chains representing OP164[cys140] and OP174[cys174] were translated in vitro in a rabbit reticulocyte translation system without the addition of membranes and subjected to AMS or QSY modification as described in the legend of Figure 6.3.


133

Figure 6. 7 AMS and QSY modification of (a) OP164[cys140] and (b) OP174[cys140], in the presence of semi-permeabilised mammalian cells. OP164[cys140] and OP174[cys174] were synthesised in a rabbit reticulocyte translation system in the presence of digitonin-permeabilised cells and subjected to AMS or QSY modification as described in the legend of Figure 6.2.


134

In the presence of the ER translocon, OP164[cys140] was shown to be modified by both

AMS and QSY (Fig. 6.7a, lanes 1 and 2, 3 and 4), and when the nascent chain was

extended to 174 residues, both AMS and QSY were still able to react with cys140 (Fig.

6.7b, lanes 1 and 2, 3 and 4). Taken together, this analysis shows that the loop region

following TM3 remains in a hydrophilic environment as it moves out of the ribosome

and confirms that the failure of AMS to modify cys124 in the OP164 intermediate is not

due to a ‘gating’ phenomenon.

6.7 The OP164 nascent chain represents a true integration intermediate that is

attached to the ribosome

The analysis of the nascent chain environment during integration by using thiol-specific

probes relies on the assumption that the ribosome bound integration intermediates are

stable and reflect the environment of the nascent chain within the ER translocon. In

order to test this assumption, an established assay was utilised to verify the association

of nascent opsin chains with the ribosome (Wilson et al., 2005). This is of paramount

importance for OP164[cys124] since it is important to establish that the lack of AMS

modification (section 6.4) is not simply a consequence of the specific release of these

opsin chains from the ribosome resulting in their release into the phospholipid bilayer

which would preclude AMS modification.

In this case, OP164[cys124] integration intermediates were synthesised in the presence

of semi-permeabilised mammalian cells. The translation reaction was then halted with

the addition of either cycloheximide, which stabilises ribosome-nascent chain

complexes, or puromycin, which releases nascent chains from the ribosome. The

membrane fraction was isolated by centrifugation and then treated with a non-ionic

detergent, C12E8, which efficiently solubilises the membranes without disrupting any

ribosome-nascent chain complexes (Wilson et al., 2005). The ribosomes and any

associated nascent chains were recovered by centrifugation, and products in the pellet

and the supernatant were subsequently resolved by SDS PAGE.


135

Figure 6. 8 Isolation of ribosomes and associated OP164[cys124] chains. OP164[cys124] was synthesised in a rabbit reticulocyte translation system in the presence of semi-permeabilised cells. The translation reaction was halted either with cycloheximide (CHX) or puromycin. The membrane fraction was isolated by centrifugation and was solubilised in 1% C12E8. An aliquot was removed and analysed on SDS PAGE as total products (T). Ribosomes and any associated nascent chains were pelleted by centrifugation and proteins in the supernatant were precipitated using trichloroacetic acid (TCA). Products in the pellet (P) and supernatant (S) were denatured in sample buffer and resolved by SDS PAGE. Doubly-glycosylated opsin chains are indicated with (ii).


136

This experiment showed that the OP164[cys124] chains were almost exclusively in the

pellet fraction when the samples were cycloheximide treated, confirming that the

nascent chains were not released from the ribosome under these conditions and were

therefore authentic integration intermediates (Fig. 6.8, lanes 1-3). On the other hand,

treatment of the nascent chains with puromycin resulted in the release of a substantial

fraction of the nascent opsin chains from the ribosome and these were found in the

supernatant fraction (Fig. 6.8, lane 6). The puromycin treatment was not completely

effective since a proportion of nascent chains was still found associated with the

ribosomes in the pellet fraction (Fig. 6.8, lane 5). Nonetheless, since the AMS and QSY

modification reactions are performed on cycloheximide treated nascent chains, it can be

concluded that the earlier observations with AMS and QSY detailed above reflect the

true environment of the opsin chain during its insertion at the ER translocon.

6.8 Summary

Distinct regions of opsin TM3 experience different environments during membrane

integration via the ER translocon, as evidenced by the use of sulphydryl specific

reagents with distinct biophysical properties. The hydrophilic loop preceding TM3, and

the N-terminal region of TM3, remain in a hydrophilic location, presumably the

aqueous environment of the ribosomal exit tunnel and the translocon pore, during

nascent chain extension from 130 to 164 residues. On the other hand, the central region

of TM3, as represented by cys124, moves from a hydrophilic to a hydrophobic

environment when the nascent chain is extended from 140 to 150 residues where it

remains at a chain length of 164 residues. This environment relates directly to the

precise segment of polypeptide within TM3, since a probe located C-terminal of TM3

(cys140) is in a hydrophilic environment.

Conclusion

Extension of the nascent opsin chain from 140 to 150 residues and beyond results in the

central region of opsin TM3 relocating from a hydrophilic to a hydrophobic

environment within the context of the ribosomal exit tunnel and/or the ER translocon.

Chapter 7 - Discussion

137

CHAPTER 7 Discussion


138

7.1 Introduction

The aim of this project was to investigate the molecular environment of the seven TM

domain protein, opsin, during its biosynthesis at the ER, so as to better understand the

mechanism by which the multiple TM domains of a polytopic protein are integrated into

the ER membrane. Opsin was chosen as a suitable model because a high resolution

structure of the fully folded protein is available (Palczewski et al., 2000). In addition,

the N-glycosylation of its N-terminal region serves as a useful marker for the correct

targeting and insertion of the truncated integration intermediates used to study its

biogenesis.

In this study, a cysteine-mediated site-specific cross-linking approach was primarily

employed to examine the environment of the TM domains of nascent opsin during

synthesis at the ER. Cross-linking using the sulphydryl specific reagent, BMH, was

carried out with integration intermediates of defined lengths to represent different stages

of nascent chain elongation. By performing immunoprecipitations using specific

antisera, molecules adjacent to the TM domain of the nascent chain at a distinct stage

during its biosynthesis could be identified, thus allowing alterations in the environment

of the TM domain to be followed. Since a heterogeneous population of nascent chains

could be generated during in vitro translation, a HA epitope tag was added to the C-

terminus of each nascent opsin chain analysed, allowing the selection of authentic

integration intermediates and their cross-linking partners by immunoprecipitation using

an α-HA antiserum. This approach facilitated the accurate interpretation of the cross-

linking data obtained during this work.

My study focused on the association of each opsin TM domain with components of the

Sec61 complex. I made the assumption that a loss of cross-linking to the Sec61α and

Sec61β subunits reflected the lateral movement of a TM domain from the protein lined

translocon and into a primarily phospholipid environment on the basis of previous

photocross-linking studies (Martoglio et al., 1995; McCormick et al., 2003; Mothes et

al., 1997). In addition, this cross-linking approach was complemented by the analysis of

TM ‘environment’ during insertion into the ER translocon using two sulphydryl specific

modification reagents with different physical properties. On the basis of the data


139

obtained during this study, a working model of the integration process for opsin from

the translocon into the ER membrane has been generated (Fig. 7.1).

Figure 7. 1 A working model for the integration of opsin into the ER membrane. Schematic representation of Sec61α and Sec61β are shown in orange, opsin TM domains are shown in blue, while PAT-10 is shown in purple. TM1 is fully engaged with the translocon when the nascent chain is ∼96 residues long. The insertion of TM2 causes TM1 to relocate to the periphery of the translocon where it associates with PAT-10 (OP130 and OP140). When the nascent chain is ∼150 residues, TM3 is now engaged with the translocon pore while TM1 re-associates with Sec61α. Extension of the nascent chain to ∼204 residues results in the insertion of TM4 into the pore and the simultaneous integration of TM1 to TM3 into the membrane. At 259 residues, TM4 exits the translocon as soon as TM5 is inserted into the pore. When the nascent chain is ∼304 residues, TM5 associates with PAT-10 but still remains close to the Sec61 complex, while TM6 is engaged with the translocon. Lengthening of the nascent chain to ∼339 and ∼357 residues allow the insertion of TM7 into the translocon.


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7.2 The integration of TM1, TM2 and TM3 is a ‘co-ordinated’ process

To analyse the environment of TM1, cross-linking was performed using a previously

characterised cysteine probe located at residue 56 (Meacock et al., 2002) using

integration intermediates ranging from 96 to 259 residues in length. This study revealed

an unexpected level of complexity in the behaviour of TM1 which had not been

apparent during the previous analysis (Meacock et al., 2002). At the shortest chain

length examined (OP96), TM1 has fully emerged from the ribosome and engaged the

Sec61 translocon. TM1 remains at the translocon at 109 residues but moves away once

the nascent chain is extended to 130 residues, only to re-associate with the translocon at

150 and 164 residues (Fig. 7.1 and 7.2). Further extension of the nascent chain beyond

164 residues then allows TM1 to exit the translocon again. The association of TM1 with

the Sec61 complex for a second time at a chain length of 150 and 164 residues

generated adducts with Sec61α, but not Sec61β, implying this re-association may occur

at a different location to that occupied when TM1 first engages the Sec61 translocon at

a nascent chain length of 96 residues, when both Sec61α and Sec61β adducts are seen.

In addition, TM1 is in close proximity to PAT-10 when the nascent chain is 150 and

164 residues. For simplicity, I have termed the first location where cross-links to both

Sec61α and Sec61β were observed as a ‘Phase I’ environment, while the second

location, where only adducts with Sec61α was seen, as ‘Phase II’ environment (denoted

as ‘I’ and ‘II’ in Fig. 7.1 and 7.2).

Interestingly, this periodicity in the interaction of TM1 with the Sec61 subunits is only

observed when the subsequent TM domains (TM2 to TM7) are present. When TM2 to

TM7 of opsin are replaced with a hydrophilic region, TM1 relocation from the phase I

Sec61 environment is significantly delayed. Unlike the behaviour of TM1 in the normal

opsin polypeptide chains, TM1 in the context of chimeric OPTM1PPL chains remains

adjacent to Sec61α and Sec61β for a prolonged period of nascent chain synthesis. Thus,

TM1 only appears to exit the translocon when the nascent chain is longer than 204

residues (Fig. 7.2). This implies that the synthesis of additional TM domains is

necessary for authentic TM1 movement out of the translocon. The simplest explanation

for this observation is that the subsequent TM domains promote TM1 exit by displacing

it from the core of the translocon. This possibility is substantiated by the observation


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that TM2 has entered a phase I-like environment when TM1 is out of the translocon

(Fig. 7.2, OP140).

Figure 7. 2 The roles of TM1 and TM3 during opsin integration. The Sec61 complex, opsin TM domains and PAT-10 are represented in orange, blue and purple respectively. See text for details.

The OP164 integration intermediate represents an important stage in opsin biosynthesis

where TM1 ‘returns’ to the Sec61 complex whilst TM2 is still associated with the

translocon and TM3 is located in the phase I environment (Fig. 7.2, OP164). Extension

of the nascent chain to 204 residues resulted in the loss of Sec61 adducts from all three

TM domains, implying the concurrent release of TM1, TM2 and TM3 from their

association with the translocon (Fig. 7.2, OP204). This suggests that, following its

initial exit, TM1 re-associates with the Sec61 complex to allow an association with

TM2 and/or TM3, resulting in the synchronised integration of TM1 to TM3 into the ER

membrane. In support of this model, previous studies of opsin point mutants indicated

that the stability of the full length protein is influenced by interactions between TM1

and TM2 (Bosch et al., 2003). Furthermore, the expression of opsin fragments in vivo

showed that stable membrane integration occurs only when at least TM1 and TM2 are

present in a single polypeptide chain (Heymann & Subramaniam, 1997). This particular

behaviour of opsin TM1 appears similar to that observed in studies of model proteins


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with two transmembrane domains where the integration of TM1 and TM2 appeared to

be co-ordinated (Heinrich & Rapoport, 2003; Sauri et al., 2005).

The observation that TM3 exit from the ER translocon is not dependent upon the

presence of later TM domains (TM4 to TM7), as evidenced by the OPTM1-3PPL

polypeptide, is also consistent with the view that TM1, TM2 and TM3 interact with one

another to form a folding domain that can independently exit the ER translocon (Fig.

7.2). This notion is consistent with an in vivo expression study in which two separate

opsin fragments, TM1-3 and TM4-7, could be co-expressed to form a ‘functional’ opsin

molecule with spectral properties similar to the wild type protein (Ridge et al., 1995).

Thus, the integration of TM1, TM2 and TM3 into the ER membrane is likely to be a

‘co-ordinated’ process where mutual stabilisation of TM1, TM2 and TM3 acts to

facilitate the lateral exit of these TM domains from the translocon. Mutual stabilisation

between TM domains during nascent chain integration into the ER membrane is not

unique to opsin and appears to contribute to the integration of other polytopic

membrane proteins such as Neurospora plasma membrane H+-ATPase and human P-

glycoprotein (Lin & Addison, 1995; Skach & Lingappa, 1993).

7.3 An alternative analysis of TM3 environment during opsin integration

In order to further define the integration of opsin TM3, monofunctional sulphydryl-

specific reagents of different hydrophilicity, AMS and QSY, were utilised to examine

the environment of single cysteine residues introduced at different locations within

TM3. The exclusion of AMS from hydrophobic environments meant that only cysteine

probes present in an aqueous surrounding should be AMS modified, while QSY should

modify cysteine residues present in both hydrophilic and hydrophobic environments.

Using this approach, I found clear evidence that the presence of the ER translocon

specifically altered the local environment of a cysteine probe located in the middle of

opsin TM3. In particular, cys124 relocated from an aqueous environment in the OP140

intermediate into a hydrophobic environment in the OP150 and OP164 intermediates.

The most likely explanation for this observation is that the cysteine probe relocates from

the water lined exit tunnel of the ribosome to a hydrophobic region of the ER translocon

as the chain gets longer. In contrast, cys115 remains accessible to AMS at each chain


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length tested and hence does not undergo a similar ‘relocation’. Why should two

different cysteine residues display different properties in the same intermediate? Two

possibilities present themselves and both impact upon how we view the membrane

insertion process.

In the first scenario, TM3 would have formed an α-helix within the ER translocon

where its orientation would be relatively fixed (c.f. McCormick et al., 2003). A helical

wheel plot predicts that cys115 and cys124 would be on different sides of such an α-

helix. Thus, one side of the TM domain could be partitioning into the hydrophobic lipid

bilayer (cys124) whilst the other remained in the more hydrophilic environment of the

Sec61 complex (cys115) (‘Partitioning’ model, Figure 7.3a).

Figure 7. 3 Possible models of TM3 integration into a hydrophobic environment. The nascent opsin chain is represented by a black solid line, while the numbers indicate the relative position of the cysteine residues. The hydrophilic and hydrophobic environments are shown in blue and brown respectively. See text for details.

The second scenario allows for the lateral gating of the Sec61 translocon to be in the

form of a ‘zipper’ (‘Zipper’ model, Figure 7.3b). In this case, the region towards the

cytosolic face of the ER membrane could open laterally and allow a portion of opsin

TM3, including cys124, to access the lipid bilayer. In contrast, in the OP164

intermediate, the region of the Sec61 complex towards the ER lumen could remain

closed thereby retaining the other part of TM3, including cys115, in a hydrophilic

environment. Thus, whilst my cross-linking data leads me to present a series of


144

‘snapshots’ representing distinct stages of membrane integration, my preliminary use of

‘probe access’ implies there is most likely a more refined and complex behaviour for

individual TM domains than the composite models that I present are capable of

expressing.

It should be noted that an alternative explanation for the differences in probe access that

I observe is that patches of lipid molecules are present within the core of the Sec61

translocon (McCormick et al., 2003) which could not be observed in the crystal

structure (Van den Berg et al., 2004). Cys124 may be buried in one of these

hydrophobic patches while cys115 remains in the aqueous translocon pore. In order to

differentiate between these different models, an AMS accessibility ‘scan’ of TM3 could

be performed using nascent chains with single cysteine residues introduced at positions

along the entire length of the TM domain.

Interestingly, the different environments of cys124 and cys115 were also reflected in

their cross-linking partners. When the nascent opsin chain is 164 residues long, cys124

formed an adduct only with Sec61β while cys115 formed adducts with both Sec61β and

Sec61α. This observation implied that there may be a correlation between Sec61α

adduct formation and AMS accessibility, although further examination of nascent

chains with varying lengths will be necessary to confirm this. It is possible that adduct

formation with Sec61α occurs only when the cysteine probe is in the aqueous pore and

not when it is in a hydrophobic environment. Although the function of Sec61β is

currently unknown, the formation of Sec61β cross-links alone may reflect an ‘early’

stage in nascent chain translocation, especially since the single cysteine residue of

Sec61β is present in its cytosolic domain. In fact, the use of small molecular weight

inhibitors which block an early stage of translocation, enhanced the formation of

Sec61β cross-links with nascent chains (Besemer et al., 2005; Garrison et al., 2005).

Thus, Sec61β may have a role in facilitating the insertion of some TM domains into a

‘phase I’ like environment within the Sec61 translocon.


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7.4 Opsin TM4 exits the translocon upon nascent chain extension

Unlike the apparently synchronised integration of TMs 1-3, the integration of opsin

TM4 into the ER membrane is a relatively straightforward process. Hence, TM4 is

inserted into the core of the ER translocon when the nascent chain is 204 residues long

and exits the translocon upon chain extension to 259 residues, at which stage TM5 is in

the translocon pore (Fig. 7.1, OP204 and OP259). TM4 is fully integrated into the lipid

bilayer by the time the nascent chain is 259 residues since further extension of the

nascent chain does not result in any detectable re-association of TM4 with translocon

components. There was a suggestion that TM4 associates with a PAT-10 like

component that was observed when cross-linking was carried out with chains of 259

residues and longer. However, the efficiency of adduct formation was low and the

significance of this cross-linking product remains to be fully established.

7.5 Opsin TM5 is engaged with the ER translocon throughout opsin biosynthesis

One of the limitations of site-specific cross-linking is that the location/orientation of the

probe may influence the efficiency of the cross-linking reaction. In the study of opsin

TM5, two single cysteine probes located within TM5 did not generate any adducts to

Sec61 components, even at the shortest chain length in which the TM domain is

expected to engage the ER translocon. This may reflect the relative positions of the

cysteine residues within TM5 or a failure of this TM to engage the Sec61 complex.

However, when a cysteine probe (cys229) located in a hydrophilic loop region four

residues from the boundary of TM5 (residues 200 to 225) was analysed, discrete

adducts to Sec61 subunits were seen. This probe was therefore used for the remainder of

the analysis on the basis that it most likely reflects the associations of the nearby TM

region. Nevertheless, further studies with probes located within the hydrophobic TM

region will be needed to fully validate the conclusions based on the data obtained using

this probe location.

A further level of complexity with regard to the analysis of TM5 is that long integration

intermediates have to be analysed, and such chains generally give low efficiency

adducts that can be difficult to unambiguously confirm as being with the authentic

intermediate (see Chapter 5). In order to simplify the interpretation of such cross-linking


146

data, a shortened opsin mutant containing only the N-terminal region and TM5 to TM7

of opsin (OPN/5-7) was generated. Since the relative position of the TM domains was

altered, i.e. TM5 is now the first TM domain and so on, the resulting data must be

interpreted with care and, as much as possible, any cross-linking data obtained using

OPN/5-7 nascent chains was compared with that from equivalent authentic opsin

intermediates.

Although TM5 is effectively the first TM domain in any OPN/5-7 nascent chains, TM5

does not behave like TM1 during opsin biosynthesis. When cross-linking patterns of

TM1 integration intermediates were compared to those of equivalent lengths of TM5

OPN/5-7 integration intermediates, differences in adduct formation to Sec61

components were observed. Whilst TM1 exhibits a periodic interaction with the Sec61

complex, TM5 in the context of OPN/5-7 chains is in constant association with Sec61

components throughout nascent chain synthesis. This suggests that, unlike TM1 to

TM4, TM5 remains associated with the translocon until the complete polypeptide chain

is synthesised (see Fig. 7.1). This is significant because it implies that the translocon is

able to distinguish between different TM domains and perhaps somehow regulate their

movement into the ER membrane. This is consistent with the observation that TM

domains from various precursors are specifically positioned within the translocon,

suggesting a sequence-dependent interaction between TM domains and the Sec61

complex (McCormick et al., 2003).

One of the most striking features of the cross-linking analysis of TM5 is the strong

adduct formation with PAT-10, akin to that observed with TM1. Although it is possible

that TM5 is cross-linked to a different ∼10 kDa molecule, this is highly unlikely since

only one adduct with a ∼10 kDa protein was observed when cross-linking was

performed with opsin chains containing two cysteine residues, one in TM1 and the other

in TM5. The formation of PAT-10 adducts with TM5 in the OPN/5-7 integration

intermediates is not simply due to the relative position of TM5 as the first TM domain

in the polypeptide chain. TM5 can still form adducts with PAT-10 when cross-linking

was performed using authentic opsin nascent chains. TM5 association with PAT-10

upon nascent chain extension suggests that TM5 may relocate with respect to the

translocon during opsin biosynthesis resulting in reduced adduct formation with Sec61β

(see Fig. 7.1).


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7.6 Opsin TM6 and TM7 are associated with the Sec61 complex throughout opsin

biosynthesis

As for the analysis of TM5, the environment of TM6 and TM7 was examined using the

shorter opsin mutant, OPN/5-7. In the context of both authentic opsin chains and

OPN/5-7 chains, TM6 was found associated with the Sec61 complex from the shortest

integration intermediate examined to the full length polypeptide chain, indicating that

TM6 engages the translocon for a prolonged period of time (Fig. 7.1). Like TM6, the

cross-linking pattern of TM7 in the context of both authentic opsin and OPN/5-7

derived chains also showed that TM7 remains in the translocon throughout nascent

chain synthesis (Fig. 7.1). Since TM6 and TM7 are near the C-terminal end of full

length opsin, it is unclear whether their prolonged association with the translocon is due

to specific interactions with the Sec61 complex, or because the polypeptide ‘tether’

from the ribosome is too short to allow TM6 and TM7 to be released from the proximity

of the translocon. Hence, although ∼49 residues are present between the

peptidyltransferase centre of the ribosome and the C-terminal boundary of TM7, up to

40 residues may be within the ribosome (Kowarik et al., 2002). One possible way of

determining whether the translocon plays an active role in retaining TM6 and TM7

would be to lengthen the C-terminal polypeptide tether and see if the additional chain

length allows TM6 and TM7 to move away from the translocon.

7.7 Nascent opsin chains engage a single copy of the Sec61 complex during opsin

biosynthesis

The crystal structure of an archeal Sec61 complex suggests that a single Sec61

heterotrimer is sufficient to form the central aqueous pore of the ER translocon (Van

den Berg et al., 2004), although low resolution EM structures indicate that the Sec61

complex is normally found in the form of an oligomer composed of three or four copies

of the heterotrimer (Beckmann et al., 1997; Hanein et al., 1996; Menetret et al., 2000;

Menetret et al., 2005). The recent observation that a cysteine residue located in the

centre of the related SecY complex could form a disulphide bond with a translocating

nascent chain supports the hypothesis that the pore of the translocon resides within a

single Sec61 complex (Cannon et al., 2005), and hence the functional relevance of the

oligomeric status of the Sec61 complexes is currently unclear. However, several


148

potential reasons have been postulated; for example, it might be that an interaction

between Sec61 complexes is necessary to ‘activate’ one heterotrimer to form the

channel of the ER translocon. Alternatively, whilst one Sec61 heterotrimer forms the

active channel, the remaining complexes of the oligomer may recruit accessory

components involved in co- or post-translocational modifications such as the SPC and

OST complexes (Dobberstein & Sinning, 2004).

Since each Sec61 complex within the oligomer may be capable of forming an active

pore, different substrates or different regions of a single nascent chain might be

simultaneously translocated/integrated at separate pores within the same higher order

translocon. In the case of a polytopic membrane protein, it was suggested that different

TM domains could be translocated concurrently into the distinct pores of adjacent

Sec61 complexes within the same translocon (Dobberstein & Sinning, 2004). Double

probe analysis was employed during this study in order to investigate this possibility. I

found that the different TM domains of a single nascent opsin chain are adjacent to only

one copy of the Sec61α subunit, and conclude that it is unlikely that the different TM

domains of a polytopic membrane protein utilise separate Sec61 heterotrimers during

their integration and that only one Sec61 complex forms the active pore of the

translocon (c.f. Cannon et al., 2005).

7.8 The possible role of PAT-10 as a TM chaperone

An interesting feature observed during the analysis of opsin integration is the adduct

formation between specific TM domains and a novel ∼10 kDa protein, PAT-10. Cross-

linking between opsin and PAT-10 was first observed during an extensive analysis of

TM1 environment (Meacock et al., 2002). However, adduct formation with PAT-10 is

not specific to opsin since similar adducts were seen with the rat neurotensin receptor

(Meacock et al., 2002). In this study, PAT-10 adducts were also observed with opsin

TM4 (weakly) and TM5 (strongly), in addition to TM1. PAT-10 cross-linking to TM1,

TM4 and TM5 is characterised by an adduct which normally appears after the

subsequent TM domain has emerged from the ribosome and remains until the full length

opsin chain has been synthesised. Since nascent chain extension is necessary for the TM

domains to form adducts with PAT-10, this suggests that TM domain relocation is

required for association with PAT-10. Proximity is lost when the nascent chain is


149

released from the ribosome, indicating that PAT-10 is most likely associated with the

translocon (Meacock et al., 2002). This idea is strongly supported by my double

cysteine probe analysis which showed that a single nascent chain could be cross-linked

to both Sec61β and PAT-10 at the same time. This analysis also indicated that, like the

Sec61 complex, only a single copy of the PAT-10 protein was adjacent to an opsin

chain during its membrane integration.

Although the identity of PAT-10 is currently unknown, PAT-10 has been postulated to

have a role as a TM chaperone (Meacock et al., 2002). It is possible that PAT-10

functions to regulate the ‘release’ of specific TM domains to allow their co-ordinated

integration into the ER membrane. For example, the interaction of PAT-10 with TM1

might facilitate its assembly with TM2 and TM3 for a synchronised membrane

integration. The novel observation that PAT-10 interacts specifically with TM1, TM4

and TM5 is interesting because these TM domains may represent the beginnings of

separate folding domains. Although distinct subregions are not obvious from the crystal

structure (Palczewski et al., 2000), in vivo expression of opsin fragments has shown that

TM1-3 and TM4-7 (Ridge et al., 1995), or TM1-4 and TM5-7 (Ridge et al., 1996), can

be correctly assembled to produce functional opsin molecules. The potential role of

PAT-10 as a TM chaperone further supports the idea that TM integration into the ER

membrane is a highly regulated event.

7.9 Conclusion

The use of opsin as a model polytopic membrane protein to study the integration of TM

domains into the ER membrane underscores the fact that the integration of multiple TM

domains is a highly complex process. The working model I have produced for opsin

integration (Fig. 7.1) does not precisely conform to either the ‘sequential’ or the ‘en

masse’ model (see Chapter 1), but rather combines elements of both models. During

integration, each TM domain of opsin behaves in a unique manner, with some TM

domains engaging the translocon for longer periods than others. Opsin TM1 displays a

complex behaviour where it initially moves away from the translocon after its insertion

only to return at a later stage, most likely to interact with TM2 and TM3 in order to

allow a co-ordinated integration of these three TM domains into the ER membrane.

TM4, on the other hand, exits the translocon in an independent manner almost as soon


150

as the next TM domain has been inserted into the translocon pore. TM5, TM6 and TM7

appear to remain associated with the Sec61 complex throughout nascent chain synthesis

and presumably integrate into the membrane once the polypeptide chain is released

from the ribosome. This study has also provided further evidence that the integration of

the multiple TM domains of a polytopic membrane protein is a highly regulated process

that involves the interplay of various components such as the Sec61 complex and the

potential chaperone PAT-10.

Chapter 8 - Bibliography

151

CHAPTER 8 Bibliography


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APPENDICES

Appendices

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(A)

Intensity of products Nascent chain Opsin band Sec61α adduct

Fraction of Sec61α adduct

Relative fraction to OP96

OP96 530932 50055 0.0943 1.00 OP109 1355772 97392 0.0718 0.76 OP130 - - - - OP150 11074 418 0.0377 0.40 OP164 722440 63663 0.0881 0.93 OP204 - - - - OP259 - - - -

(B)

Intensity of products Nascent chain Opsin band Sec61α adduct

Fraction of Sec61α adduct

Relative fraction to OPTM1PPL164

OPTM1PPL109 16493.9 824.4 0.0500 0.94 OPTM1PPL130 26036.5 898.6 0.0345 0.65 OPTM1PPL150 71729 2870.7 0.0400 0.75 OPTM1PPL164 47964.4 2553.9 0.0532 1.00 OPTM1PPL204 86374.9 3942.5 0.0456 0.86 OPTM1PPL259 - - - -

Appendix 1.1 Raw data for the quantification of products obtained from the cross-linking analysis of integration intermediates of (A) normal opsin and (B) OPTM1PPL polypeptide chain. Where appropriate, products due to uncross-linked doubly-glycosylated opsin chains and Sec61α adducts from immunoprecipitations using the α-opsin antibody were quantified using the AIDA software. The fraction of Sec61α adduct was obtained by dividing the intensity of the Sec61α adduct by the value obtained for doubly-glycosylated opsin chains. The integration intermediate with the highest fraction was set to a nominal value of 1.00 and the relative fractions of Sec61α adduct formation for the other intermediates were calculated.

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173

M N G T E G P N F Y V P F S N K T G V V R S P F E A P Q 28

Y Y L A E P W Q F S M L A A Y M F L L I M L G F P I N F 56

L T L Y V T V Q H K K L R T P L N Y I L L N L A V A D L 84

F M V F G G F T T T L Y T S L H G Y F V F G P T G C N L 112

E G F F A T L G G E I A L W S L V V L A I E R Y V V V C 140

K P M S N F R F G E N H A I M G V A F T W V M A L A C A 168

A P P L V G W S R Y I P E G M Q C S C G I D Y Y T P H E 196

E T N N E S F V I Y M F V V H F I I P L I V I F F C Y G 224

Q L V F T V K E A A A Q Q Q E S A T T Q K A E K E V T R 252

M V I I M V I A F L I C W L P Y A G V A F Y I F T H Q G 280

S D F G P I F M T I P A F F A K T S A V Y N P V I Y I M 308

M N K Q F R N C M V T T L C C G K N P L G D D E A S T T 336

V S K T E T S Q V A P A 348

Appendix 1.2 The amino acid sequence of bovine opsin. Cysteine residues which have been replaced with glycine residues to obtain the cysteine null opsin polypeptide chain are in blue, while residues which have been converted to cysteine residues for site-specific cross-linking are in red. Regions representing transmembrane domains are underlined in black.


Y Y L A E P W H E E T N N E S F V I Y M F V V H F I I P 56

L I V I F F C Y G Q L V F T V K E A A A Q Q Q E S A T T 84

Q K A E K E V T R M V I I M V I A F L I C W L P Y A G V 112

A F Y I F T H Q G S D F G P I F M T I P A F F A K T S A 140

V Y N P V I Y I M M N K Q F R N C M V T T L C C G K N P 168

L G D D E A S T T V S K T E T S Q V A P A 189

Appendix 1.3 The amino acid sequence of the OPN/5-7 polypeptide chain. Residues 36 to 194 have been deleted from the sequence of opsin to form OPN/5-7. The position of the deleted region is indicated with ‘ ’. All other notations are as described in appendix 1.2.

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174


Y Y L A E P W Q F S M L A A Y M F L L I M L G F P I N C 56

L T L Y V T V Q H K K L R T T P V G P N G P G N G Q V S 84

L R D L F D R A V M V S H Y I H D L S S E M F N E F D K 112

R Y A Q G K G F I T M A L N S G H T S S L P T P E D K E 140

Q A Q Q T H H E V L M S L I L G L L R S W N D P L Y H L 168

V T E V R G M K G A P D A I L S R A I E I E E E N K R L 196

L E G M E M I F G Q V I P G A K E T E P Y P V W S G L P 224

S L Q T K D E D A R Y S A F Y N L L H G L R R D S S K I 252

D T Y L K L L N C R I I Y N N N C 269

Appendix 1.4 The amino acid sequence of OPTM1PPL[cys56]. Residues representing opsin TM1 is underlined in black while residues in the preprolactin sequence are shaded in grey. The cysteine probe at position 56 is in bold.


Y Y L A E P W Q F S M L A A Y M F L L I M L G F P I N G 56

L T L Y V T V Q H K K L R T P L N Y I L L N L A V A D L 84

F M V F G G F T T T L Y T S L H G Y F V F G P T G G N L 112

E G C F A T L G G E I A L W S L V V L A I E R Y V V V G 140

K P T P V G P N G P G N G Q V S L R D L F D R A V M V S 168

H Y I H D L S S E M F N E F D K R Y A Q G K G F I T M A 196

L N S G H T S S L P T P E D K E Q A Q Q T H H E V L M S 224

L I L G L L R S W N D P L Y H L V T E V R G M K G A P D 252

A I L S R A I E I E E E N K R L L E G M E M I F G Q V I 280

P G A K E T E P Y P V W S G L P S L Q T K D E D A R Y S 308

A F Y N L L H G L R R D S S K I D T Y L K L L N C R I I 336

Y N N N C 341

Appendix 1.5 The amino sequence of OPTM1-3PPL[cys115]. The transmembrane domains are underlined in black while residues due to preprolactin are shaded in grey. Cys115 is in bold.

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Appendix 1.6 Schematic representation of the secondary structure of bovine opsin (adapted from Palczewski et al., 2000). Cysteine residues which were replaced with glycines are shown in blue while residues mutated to cysteines are shown in red. Asparagine-linked glycan groups are indicated by ‘Y’ and the disulphide bridge is represented with a blue dashed line.

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