REPAIR OF CFTR DEFECTS CAUSED BY CYSTIC

FIBROSIS MUTATIONS

Li Shi

A thesis submitted in conformity with the requirements for the degree of

Master of Science, Institute of Medical Science,

in the University of Toronto

© Copyright by Li Shi 2013

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Li Shi

Master of Science, 2013

Institute of Medical Science, University of Toronto

ABSTRACT

Cystic fibrosis is caused primarily by deletion of Phe508. An exciting discovery was that

CFTR‟s sister protein, the P-glycoprotein (P-gp) containing the equivalent mutation (ΔY490),

could be repaired by a drug-rescue approach. Drug substrates showed specificity, and their

mechanism involves direct binding to the transmembrane domains (TMDs) since arginine

suppressor mutations were identified in TMDs that mimicked drug-rescue to promote maturation.

We tested the possibility of rescuing CFTR processing mutants with a drug-rescue approach. 1)

Arginine mutagenesis was performed on TM6, 8, and 12. 2) Correctors were tested for

specificity. 3) Truncation mutants were used to map the VX-809 rescue site. Correctors 5a, 5c,

and VX-809 were specific for CFTR. VX-809 appeared to specifically rescue CFTR by

stabilizing TMD1. Therefore, the TMDs are potential targets to rescue CFTR. Rescue of P-gp

and CFTR appeared to occur by different mechanisms since no arginine suppressor mutations

were identified in CFTR.

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ACKNOWLEDGEMENTS

I would like to express the deepest appreciation to Dr. David M. Clarke and Dr. Tip W.

Loo, my enthusiastic supervisors for their patience, guidance, encouragement, and advice. I

cannot say thank you enough for their continuous support of my Master‟s study and research.

Furthermore, I would like to thank the technician in the lab, Claire M. Bartlett, for teaching me

all the lab techniques used to produce this thesis as well for the support on the way. It would not

have been possible to finish this thesis without the guidance of my committee members, Dr.

David B. Williams and Dr. Walid Houry. I would like to thank them for taking the time to offer

their advice and ask me hard questions to keep me thinking along the way. Finally, my most

sincere thanks to my parents for their unconditional support, both financially and spiritually

throughout my degree.

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CONTRIBUTIONS

Dr. Tip W. Loo: All mutants used in this thesis were constructed by Dr. Tip W. Loo. (See section

2.1 (Construction of Mutants))

Claire M. Bartlett: All methods in Section 2 of this thesis were taught to me by Claire M. Bartlett.

Furthermore, she was responsible for preparing the media (DMEM) required for cell culture and

the TBS stock solution used in Western blotting.

Sections 3.2.1 and 3.2.3 of this thesis constituted a publication in Loo, T.W., Bartlett, M.C., Shi,

L., and Clarke, D.M. (2012) Corrector-mediated rescue of misprocessed CFTR mutants can be

reduced by the P-glycoprotein drug pump. Biochem. Pharmacol. 83: 345-354.

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TABLE OF CONTENTS

1 INTRODUCTION ............................................................................................................... 1

1.1 Cystic fibrosis and the CFTR gene ................................................................................... 1

1.2 Physiological role of the CFTR protein ............................................................................. 2

1.3 Structure of the CFTR protein .......................................................................................... 7

1.4 Gating mechanism of the CFTR channel........................................................................... 12

1.5 Biosynthesis and degradation of the CFTR protein ........................................................... 15

1.6 CFTR gene mutations and their consequences at the cellular level .................................... 18

1.7 Clinical manifestations and diagnosis of cystic fibrosis ..................................................... 22

1.8 Treating the basic defect of cystic fibrosis ........................................................................ 25

1.8.1 Gene therapy .......................................................................................................... 26

1.8.2 Indirect rescue approaches ...................................................................................... 28

1.8.3 Direct rescue and the use of pharmacological chaperones ....................................... 30

1.9 Experimental evidence for possibility of direct rescue ...................................................... 32

1.10 Objectives ...................................................................................................................... 36

1.10.1 Arginine scanning mutagenesis of the transmembrane segments of CFTR ............ 36

1.10.2 Direct rescue of CFTR processing mutants using correctors ................................. 38

2 METHODS .......................................................................................................................... 40

2.1 Construction of mutants .................................................................................................... 40

2.2 Cell culture ....................................................................................................................... 41

2.3 Cell surface labeling ......................................................................................................... 43

2.4 Cycloheximide chase assay .............................................................................................. 43

2.5 Western blotting ............................................................................................................... 45

2.6 Iodide efflux assay............................................................................................................ 46

3 RESULTS ............................................................................................................................ 48

3.1 Arginine suppressor mutations.......................................................................................... 48

3.1.1 Mapping the structure of CFTR TMDs and testing whether arginines introduced

in the TMDs of wt-CFTR promote maturation ........................................................ 49

3.1.2 Performing iodide efflux assays to examine mutant channel function ...................... 56

3.1.3 Identifying suppressor mutations in the TMDs of CFTR ......................................... 59

3.2 Direct rescue using correctors ........................................................................................... 63

3.2.1 Identifying correctors that specifically interact with CFTR processing mutants ....... 63

3.2.2 Identifying sites of corrector interactions ................................................................ 71

3.2.3 Effect of other mutations on stability of CFTR ........................................................ 83

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4 DISCUSSION ...................................................................................................................... 85

4.1 Arginine suppressor mutations ......................................................................................... 85

4.1.1 Conclusions ............................................................................................................ 88

4.2 Direct rescue using correctors ........................................................................................... 89

4.2.1 Conclusions ............................................................................................................. 96

4.3 Future Directions .............................................................................................................. 96

5 REFERENCES .................................................................................................................... 98

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LIST OF TABLES

Table 1 Classes of CFTR Mutations that cause cystic fibrosis. ........................................ 19

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LIST OF FIGURES

Figure 1 Schematic model of CFTR ................................................................................ 9

Figure 2 Models of CFTR and P-glycoprotein ................................................................ 35

Figure 3 Effect of arginine mutations on maturation of CFTR......................................... 51

Figure 4 Iodide efflux activity of TM6, TM8, and TM12 CFTR mutants ........................ 58

Figure 5 Model of CFTR with the locations of V232, H1085, and F508 highlighted ......... 60

Figure 6 Immunoblot analysis of the double mutants generated to test for suppressor

mutations .......................................................................................................... 62

Figure 7 Structure of correctors ....................................................................................... 65

Figure 8 Effect of correctors on H1085R CFTR and G268V P-gp ................................... 67

Figure 9 Stability of ΔF508 CFTR in the presence or absence of correctors .................... 68

Figure 10 Effect of corr-5a on expression of ΔF508 CFTR on the cell surface .................. 70

Figure 11 Effect of VX-809 on glycosylation of TMD1+2 CFTR ..................................... 73

Figure 12 Effect of VX-809 on maturation of ΔNBD2(Δ1197-1480) CFTR ...................... 75

Figure 13 Coexpression of C-half and N-half CFTRs ........................................................ 77

Figure 14 Coexpression of C-half CFTR and N-half CFTRs containing truncated NBD1 .. 79

Figure 15 Effects of VX-809 on TMD1, TMD2, and NBD1 CFTRs .................................. 80

Figure 16 Effect of VX-809 on TMD1 CFTR turnover ..................................................... 82

Figure 17 Stability of other CFTR mutants ........................................................................ 84

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LIST OF ABBREVIATIONS

ABC ATP-binding cassette

ASL airway surface liquid

BES N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid

BHK baby hamster kidney

cAMP adenosine 3‟,5‟-cyclic monophosphate

CF cystic fibrosis

CFTR cystic fibrosis transmembrane conductance regulator

CHO Chinese hamster overy

DIDS 4,4‟- Diisothiocyano-2,2‟-stilbenedisulfonic acid

DMEM Dulbecco‟s modified Eagle‟s media

DMSO Dimethyl sulfoxide

EDTA ethylenediaminetetraacetic acid

ENaC epithelial sodium channel

ER endoplasmic reticulum

ERAD ER-associated degradation

GABA γ-aminobutyric acid

Hdj-2 Human DnaJ 2

HEK human embryonic kidney

HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

Hsp70 70kDa heat shock protein

ICL intracellular loop

MDR1 Multidrug resistance protein 1

Mg-AMP-PNP Mg-adenylylimidodiphosphate

MTS methanethiosulfonate

NBD nucleotide-binding domain

OST complex oligosaccharyltransferase complex

PAGE polyacrylamide gel electrophoresis

PBS phosphate buffered saline

PBSCM PBS containing 0.1mM CaCl2 and 1mM MgCl2

P-gp P-glycoprotein

PKA protein kinase A

RFLPs restriction fragment-length polymorphisms

SDS sodium dodecyl sulphate

TBS Tris-buffered saline

TBST TBS containing 0.5% (v/v) Tween-20

TM transmembrane segment

TMD transmembrane domain

Tris tris(hydroxymethyl)-aminoethane

Triton X-100 octyl phenoxy polyethoxyethanol

1

1 INTRODUCTION

1.1 CYSTIC FIBROSIS AND THE CFTR GENE

Cystic fibrosis (CF) is the most common fatal autosomal recessive genetic disorder in

the Caucasian population – one in 2500 babies is born with the disease, and one in 25 babies

carries the CF gene (Reviewed by Rowe, 2005). As other recessive genetic disorder, only

individuals with a defective mutation in both alleles show the symptoms of the disease. The

discovery of cystic fibrosis can be dated back to the Middle Ages, when people had the

saying: “Woe to that child which when kissed on the forehead tastes salty. He is bewitched

and soon must die.” This is one of the earliest references to cystic fibrosis, recognizing the

association between the salt loss in CF and illness, although the condition was unnamed at

that time (Reviewed by Welsh and Smith, 1995). Later in 1936, the Swiss pediatrician Guido

Fanconi (1936) named the disease “cystic fibrosis with bronchiectasis”, and published a

paper describing the relationship between cystic fibrosis, celiac disease, and bronchiectasis.

In 1938, Dr. Dorothy Andersen (1938) for the first time coined the term “cystic

fibrosis of the pancreas” and provided a clear detailed clinical and pathological description of

it. She examined 49 patients and identified the major symptoms common to CF patients,

including neonatal intestinal obstruction, and histological changes in the lungs, intestine, and

pancreas – particularly the fluid-filled sacs and scar tissue observed in the pancreases of

patients. Andersen continued her research on cystic fibrosis, and in 1946, Andersen and her

colleague R.G. Hodges, through the genetic analysis of 113 families, identified the disease as

a monogenetic classic Mendelian disease that is inherited in an autosomal recessive pattern

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(Andersen and Hodges, 1946).

Knowlton et al. (1985) were able to use restriction fragment-length polymorphisms

(RFLPs) as genetic markers to localize the CF gene to chromosome 7 in 1985. Later in 1989,

research teams headed by Professor Lap-Chee Tsui, Dr. Francis Collins, and Professor Jack

Riordan identified the specific gene sequence responsible for cystic fibrosis (Riordan et al.,

1989). Using positional cloning and the techniques of chromosome “jumping” and “walking”,

they discovered that cystic fibrosis is caused by mutations in the CF gene, which contains 27

exons spreading over 250 kb of chromosome 7 (7q31.2). It encodes a protein that is 1480

amino acids long and has a molecular weight of 168,173 Da called the cystic fibrosis

transmembrane conductance regulator (CFTR). Furthermore, as supporting evidence for the

role of the CF gene in cystic fibrosis, mutational analysis was conducted to show a 3-base

pair deletion absent in normal chromosomes but present in approximately 70% of CF

chromosomes examined. The mutation discovered was the most common mutation in CF,

named ΔF508, which resulted from the deletion of the nucleotide triplet CTT in exon 10 of

the CF gene.

1.2 PHYSIOLOGICAL ROLE OF THE CFTR PROTEIN

When the CF gene encoding CFTR was discovered in 1989, it was unclear how the

protein functioned to regulate ion conductance across the apical membrane of epithelial cells.

The observation that 10 of the 12 putative transmembrane regions of CFTR contained one or

more amino acids with charged side chains suggested it to be an ion channel itself, as this

amphipathic nature of the transmembrane segments was believed to contribute to the

channel-forming capacity of the brain sodium channel and the γ-aminobutyric acid (GABA)

3

receptor chloride channel subunits (Riordan et al., 1989). However, the primary sequence of

CFTR did not resemble that of the purified polypeptides that were capable of reconstituting

chloride channels in lipid membranes, and thus, it was also suggested that CFTR could be a

chloride channel regulator rather than a chloride channel itself (Riordan et al., 1989). In 1991,

Anderson et al. (1991) demonstrated that CFTR forms a cAMP-regulated anion pore through

the studies of recombinant CFTR. They mutated basic amino acids in the putative

transmembrane domains of CFTR, and found that mutation of lysines at positions 95 or 335

to acidic amino acids altered the sequence of anion selectivity of cAMP-regulated channels

in cells containing either endogenous or recombinant CFTR. To provide further evidence that

CFTR is an apical membrane chloride channel, Anderson et al.(1991) expressed ΔF508

CFTR or wildtype CFTR in HeLa, Chinese hamster overy (CHO), and NIH 3T3 fibroblast

cells, and measured anion permeability using a fluorescence microscopic assay and the

whole-cell patch-clamp technique. It was observed that only expression of wildtype CFTR

generated a unique chloride current upon cAMP stimulation. Bear et al. (1992) have also

tested the postulate that CFTR is a regulated low-conductance chloride channel by

incorporating highly purified recombinant CFTR into planar lipid bilayers, and showing that

they form chloride channels with properties similar to those observed in native intact

epithelial cells. After numerous studies with recombinant CFTR, it is now known that CFTR

is an adenosine 3‟,5‟-cyclic monophosphate (cAMP)-regulated chloride channel located

primarily at the apical surfaces of epithelial cells in multiple tissues including the liver,

pancreas, intestine, sweat glands, and lungs, where it plays an important role in determining

transepithelial salt transport, fluid flow, and ion concentrations. In CF patients, normal

hydration of the epithelial surfaces is disrupted due to the lack of chloride channel activity of

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mutant CFTRs (Gadsby et al., 2006). Therefore, CF patients experience mucus accumulation

in a variety of ducts within organs such as the pancreas, salivary glands, sweat glands, and

lungs (Reviewed by Rowe, 2005).

A high level of sodium chloride in the sweat is a hallmark of cystic fibrosis. Sweat

test involving the measurement of the concentration of chloride ion in a sample of

pharmacologically induced sweat is the most efficient, expedient test commonly conducted

to diagnose cystic fibrosis (Reviewed by Quinton, 2007). In the sweat gland, CFTR is

involved in fluid and electrolyte secretion when expressed in the apical membrane of the

secretory coil. It participates in fluid and electrolyte absorption when expressed in the apical

and basolateral membranes of absorptive duct cells. Reddy et al. (1999) explored the role

CFTR plays in regulating the epithelial sodium channel (ENaC) activity in native human

sweat duct by calculating CFTR Cl- conductance and ENaC Na+ conductance from

transepithelial electrical conductances measured before and after stimulating CFTR with

cAMP or cGMP or GTP-γ-S in the presence of ATP. Their findings suggested that CFTR

and ENaC at the apical membrane of sweat gland ducts are activated simultaneously.

Furthermore, ENaC activation depends on CFTR function, as activation of ENaC by cAMP,

GMP, or G-proteins was not observed when Cl- was removed from the medium and when

CFTR was blocked with the inhibitor DIDS (4,4‟- Diisothiocyano-2,2‟-stilbenedisulfonic

acid). The transepithelial chloride conductance in normal human sweat duct is absent in CF

sweat duct, as demonstrated through transepithelial electrophysiological studies done by

Bijman et al. (1986). As a result, ENaC Na+ conductance is limited, and NaCl is poorly

absorbed in the CF duct. This leads to the production of sweat containing a high level of salt.

In contrast to its role in the sweat gland duct, CFTR expressed in the pancreas plays a

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key role in fluid and electrolyte secretion. Marino et al. (1991) conducted

immunocytochemical studies and localized CFTR to the apical membrane of the proximal

duct epithelial cells within the pancreas. The major task of pancreatic duct epithelial cells is

to secrete water and bicarbonate ions (HCO3- ) to neutralize the acidity of the chyme entering

the small intestine from the stomach. CFTR is involved in HCO3- transport in two ways –it

regulates the Cl-/HCO3- exchanger, and it transports HCO3

- directly. Ishiguro et al. (2009)

investigated how much of the HCO3- secretion that occurs at the apical membrane of guinea

pig pancreatic duct cells under physiological conditions is accounted for by the direct

transport of HCO3- by CFTR. They blocked HCO3

- transport via other pathways using

pharmacological agents, and used change in intracellular pH measurement to assess HCO3-

movement through CFTR. They found that the cAMP stimulated HCO3- transport was

independent of the presence of Cl- and luminal Na+, and that it was significantly inhibited

when CFTRinh-172 was used to block channel opening of CFTR. These observations

suggested that CFTR‟s major role at the apical membrane of pancreatic duct cells is to

provide a direct pathway for HCO3-. However, patch clamp studies conducted by Tang et al.

(2009) on excised membrane patches from cells heterologously expressing CF-associated

CFTR mutants showed that there was no change in HCO3- permeability in any of the three

mutants examined, suggesting that pancreatic disease in CF patients is the result of

dysregulation of the Cl-/HCO3- exchanger by CFTR. More studies need to be done to

determine the relative importance of direct and indirect HCO3- transport by CFTR.

Nevertheless, it is certain that in CF patients, a loss of CFTR function in the pancreas leads

to mucus accumulation as a result of defective secretion of bicarbonate ions. Mucus

accumulation prevents the release of digestive enzymes into the small intestine from the

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exocrine acinar cells of the pancreas, resulting in malabsorption of essential nutrients

(Reviewed by Davis, 2006). Furthermore, obstruction of the pancreatic ducts due to a build-

up of mucus can ultimately lead to atrophy and fibrosis of the pancreas, which in turn leads

to CF-related diabetes mellitus resulting from the development of endocrine pancreatic

dysfunction (Reviewed by Wilschanski et al., 2007). Fortunately, pancreatic enzyme

supplements and insulin can be given to CF patients to help them overcome CF pancreatic

dysfunction.

Most of the morbidity and mortality associated with CF today is caused by the

presence of thick tenacious secretions that obstruct distal airways and submucosal glands in

the lung. The surface liquid coating airway epithelial cells termed the airway surface liquid

(ASL) consists of two layers – a gel-like mucus layer generated by secreted mucins at the top,

and a poly-anionic watery layer known as the periciliary liquid at the bottom. In the normal

lung, apical membrane epithelial CFTR and calcium activated chloride channels work in

conjunction with the epithelial sodium channel (ENaC) to keep the height of the periciliary

liquid at 7µm, which enables efficient ciliary beating and movement of the mucus layer

towards the throat to take place (Tarran et al., 2001). Konig et al. (2001) have conducted

ENaC expression studies to demonstrate that coexpression of ENaC with either CFTR or the

ClC-0 chloride channel reduces amiloride-sensitive Na+ conductance in the presence of high

extracellular Cl-. Their data suggested that chloride currents are inhibiting ENaC in epithelial

cells. Since CFTR is the predominant chloride channel in the airways, sodium absorption is

higher in CF airways compared to non-CF airways. Therefore, in CF airways,

malfunctioning CFTR results in impaired chloride transport and sodium hyperabsorption. In

a study done by Folkesson et al. (1996), osmotic water permeability of the airways of the

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lungs was measured by dissecting and perfusing small airways from guinea pig lung with

solutions containing a membrane impermeable fluorophore called fluorescein sulfonate.

When the perfused segment is bathed in solutions of specific osmolalities, the change of

fluorescein sulfonate fluorescence resulting from transepithelial water transport is measured

and used as a measure of the osmotic water permeability of the airway. The results from their

study suggested that osmotic water permeability of the airways is high and there are water

channels present to facilitate transepithelial water movement. This implies that the

transepithelial solute concentration gradient is kept small in the airways, and that the

abnormal respiratory epithelial NaCl transport in cystic fibrosis would result in a decrease of

ASL volume. This is indeed what was observed in an in vivo study performed by Mall et al.

(2004), where they generated mice with airway-specific overexpression of ENaC to show

that sodium hyperabsorption causes ASL depletion. When the airway surface fluid is

depleted, mucociliary clearance collapses and ultimately airway obstruction due to mucus

accumulation, inflammation, repeated infections, and bronchiectasis leads to a decline in

respiratory function and eventually to lung failure (Reviewed by Rowe, 2005).

1.3 STRUCTURE OF THE CFTR PROTEIN

Riordan et al. (1989) examined the sequence of CFTR and compared it to sequences

of P-glycoprotein and other members of the ATP-binding cassette (ABC) transporter family

to determine its structure. They proposed that CFTR belongs to the ABC superfamily of

transporter proteins. It is composed of two repeated motifs linked by a unique highly charged

cytoplasmic domain containing multiple consensus phosphorylation sites that is not present

in other ABC transporters called the R domain. Each of the two repeated motifs contains a

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transmembrane domain (TMD) and a nucleotide-binding domain (NBD) (Figure 1).

Serohijos et al. (2008) confirmed the proposed structure of CFTR by constructing a

homology model using the crystal structure of bacterial multidrug ABC transporter Sav1866

as a template. In addition, Rosenberg et al. (2011) recently studied CFTR structure by

electron crystallography. They crystallized CFTR in the outward facing state and confirmed

its resemblance with the Sav1866 transporter. The two TMDs form a low conductance anion-

selective pore containing a deep and wide intracellular vestibule, and a shallow extracellular

vestibule separated by a selectivity filter located at the narrowest region of the pore

(Reviewed by Hwang and Sheppard, 2009). They are an essential part of the channel pore of

CFTR, and are responsible for conductance and selectivity of the channel pore. Mutagenesis

studies done by Anderson et al. (1991) have demonstrated that wildtype ion selectivity was

changed from Br->Cl->I->F- to I->Br->Cl-> F- when certain positively charged lysine residues

in TMD1 were mutated to negatively charged aspartic acid or glutamic acid. Bai et al. (2010)

provided further evidence that the sixth transmembrane segment (TM6) of the CFTR channel

governs the gating and conductance of the channel pore. They performed cysteine-scanning

mutagenesis in TM6 and identified charged residues that function to attract anions into the

outer mouth of the channel pore. Mutating any one of these residues to cysteine had a

negative effect on the single-channel current amplitude. Moreover, application of the

positively charged 2-trimethylammonium-ethyl MTS (MTSET) to some of the cysteine

substituted residues in TM6 altered the open time and opening rate of the channel, suggesting

that TM6 governs the channel gating behavior. Investigation of TMD2 of CFTR done by

Cotten et al. (1996) has revealed that the fourth intracellular loop (ICL4) between TM10 and

TM11 of CFTR also plays a role in the gating of the channel pore. In particular, when the

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Figure 1 Schematic model of CFTR. CFTR contains two transmembrane domains – TMD1

and TMD2 shown in light green and dark green respectively, two nucleotide-binding

domains – NBD1 and NBD2 shown in light pink and dark pink respectively, and a regulatory

domain (R). Phenylalanine 508, shown in yellow, is a residue critical for NBD1 and TMD2

interactions, as it is predicted to mediate the interaction between NBD1 and the fourth

intracellular loop – the loop connecting transmembrane segments 10 and 11 in TMD2. The

structure was generated and viewed using the PyMOL Molecular Graphics System (DeLano,

2002), which is based on the theoretical model of CFTR structure proposed by Serohijos et al.

(2008).

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residue R1066 was mutated to a cysteine, the open time of the channel was shortened, and

when the same residue was mutated to a histidine, the open probability of the channel was

increased.

The R domain of CFTR contains multiple consensus phosphorylation sites – eight

serines and one threonine residues. Regulation of CFTR channel activity is achieved by

phosphorylation of the R domain by kinases, particularly protein kinase A (PKA). However,

exactly how the R domain functions to regulate channel activity is not clear. On one hand, it

was proposed by some that the unphosphorylated R domain acts as an inhibitor that prevents

the channel from opening, and this inhibition is relieved upon phosphorylation. Evidence

supporting this idea comes from studies conducted by Rich et al. (1991), in which a CFTR

mutant lacking amino acid residues 708 – 835 from the R domain was expressed in HeLa

cells. Whole cell and SPQ fluorescence showed that the deletion produced channels that were

constitutively active – the channels opened in the presence of ATP even without

phosphorylation, suggesting that the R domain, or at least a portion of the R domain, inhibits

the constitutive activity of the channel by keeping the channel closed while in the non-

phosphorylated state. Also supporting an inhibitory role of the R domain was the observation

that mutating all of the phosphorylatable serines in the R domain significantly reduces the

channel activity (Cheng et al., 1991). Therefore, phosphorylation of the R domain eliminates

its inhibitory effect on the channel. On the other hand, it was proposed by others that the

phosphorylated R domain stimulates channel activity. Studies done by Winter et al. (1997)

have demonstrated that CFTR variants with the R domain deleted, which was able to open in

the presence of ATP without phosphorylation, displayed significantly lower open probability

than phosphorylated wildtype CFTR Cl- channels. Furthermore, they proposed that

11

phosphorylation of the R domain stimulates channel activity by enhancing the interaction of

ATP with the NBDs. Kinetic analyses conducted by Li et al. (1996) have also suggested that

PKA phosphorylation of the R domain increases the affinity of the NBDs for ATP to

enhance CFTR ATPase activity, and thereby stimulates channel activity. The mechanism by

which the R domain regulates channel activity is not known. It seems that it exerts both

inhibitory and stimulatory effects. Nevertheless, the R domain plays an important role in

regulating channel activity.

Activation of the CFTR channel requires not only the phosphorylation of the R

domain by kinases, but also the binding and hydrolysis of ATP by the NBDs. Recently, the

crystal structures for isolated mouse and human NBD1 and NBD2 have been determined

(Lewis et al., 2003, 2005; Zhao et al., 2008). CFTR‟s two NBDs are structurally asymmetric,

with only 27% amino acid identity between the two. They both contain conserved Walker A

and Walker B motifs which are essential for ATP binding and hydrolysis (Reviewed by

Gadsby et al., 2006). Structural and functional studies of other ABC transporters and

ATPases have revealed that the Walker A motif contains a lysine residue that makes direct

contact with either the α- or the γ-phosphate of ATP, the Walker B motif contains a aspartate

residue which coordinates the catalytic Mg2+ essential for ATP hydrolysis, and the highly

conserved LSGGQ motif found between the Walker A and B motifs in NBD1 is responsible

for coupling the energy from ATP hydrolysis to channel gating by direct interaction with the

transmembrane domains (Reviewed by Sheppard and Welsh, 1999). Homology modeling

conducted by Lewis et al. (2004) has revealed that in the open state of the channel, the NBDs

form a head-to-tail dimer with the two ATP-binding sites buried at the interface of the dimer

– one site formed by the Walker A and B motifs of NBD1 and the LSGGQ motif of NBD2,

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and the other formed by the Walker A and B motifs of NBD2 and the LSGGQ motif of

NBD1. This model is consistent with recently solved high resolution full length crystal

structures and NBD structures of other ABC transporters (Locher et al., 2002; Hollenstein et

al., 2007; Pinkett et al., 2007; Dawson and Locher; 2006).

1.4 GATING MECHANISM OF THE CFTR CHANNEL

The gating of the CFTR channel is mediated by both phosphorylation of the R domain

and binding and hydrolysis of ATP at the NBD domains (Vergani et al., 2003). Elevation of

cAMP level leads to activation of cAMP-dependent protein kinase A, which is capable of

phosphorylating the phosphorylatable serines in the R domain. Once the R domain is

phosphorylated, ATP binds to the NBDs to cause dimerization of the NBDs, which in turn

leads to opening of the channel. When ATP is hydrolyzed, the NBDs disassociate and the

channel closes (Vergani et al., 2003).

ATP binding at both NBDs is required for opening of the channel to occur. Using

patch clamp techniques, Vergani et al. (2003) have shown that introducing mutations into the

conserved Walker motifs of either NBD1 (K464A) or NBD2 (D1370N, K1250A) caused the

mutant CFTR channels expressed in Xenopus oocytes to open less frequently at low Mg-ATP

concentrations. Moreover, they observed that the opening rates of the mutant CFTR channels

can be restored to normal by increasing the Mg-ATP concentration – the opening rates of

K464A, D1370N, and K1250A CFTR channels were comparable to that of wildtype CFTR

channel at saturating Mg-ATP concentration. Structural information and nucleotide

photolabeling data have suggested that K464A, D1370N, and K1250A mutations reduced the

apparent affinity of the ATP binding sites (Vergani et al., 2003). Therefore, the observation

13

that defects in channel opening caused by mutations can be restored by increasing Mg-ATP

concentration suggests that ATP binding at both NBD1 and NBD2 is required for a CFTR

channel to open. Further evidence supporting this idea comes from studies conducted by

Berger et al.(2005), in which non-conserved positions of each NBD Walker A motif were

mutated by site-directed mutagenesis to phenylalanine to sterically block ATP binding. The

observation that phenylalanine substitutions in the Walker A motif of each NBD blocked [α-

32P]8-N3-ATP labeling of the mutated NBD and reduced channel opening rate suggests that

normal channel opening requires ATP binding to both NBDs.

While CFTR channel opening requires ATP binding, ATP hydrolysis is required for

closure of the CFTR channel. Evidence supporting this idea comes from the observation that

channel open time is prolonged when the ATP hydrolysis cycle is arrested by adding Mg-

adenylylimidodiphosphate (Mg-AMP-PNP), a non-hydrolyzable analogue of Mg-ATP, or

orthovanadate (VO4), an ATPase inhibitor, to the Mg-ATP used to activate the channels

(Gunderson and Kopito, 1994). Further studies have suggested that the ATP binding site of

NBD1 is catalytically inactive, and channel closing is catalyzed by ATP hydrolysis at the

NBD2 site. The crystal structure of NBD1 solved by Lewis et al. (2004) confirmed the

catalytically inactive site of NBD1. They found that the ATP binding site of NBD1 lacks the

crucial Walker B glutamate residue that serves as a catalytic base for ATP hydrolysis in

active ABC transporters. Vergani et al. (2003) also presented results suggesting closing of

CFTR channels is linked to ATP hydrolysis at NBD2. They analyzed the gating kinetics of

CFTR channels mutated at key catalytic site residues in either NBD1 or NBD2, and found

that NBD1 mutations did not significantly alter the mean channel closing rate, whereas

NBD2 mutations dramatically slowed channel closing. Therefore, it can be concluded that

14

normal rapid closing of CFTR channels is preceded by ATP hydrolysis at NBD2.

It is generally accepted that there is a strict coupling between the ATP hydrolysis

cycle and the gating cycle of CFTR, however, how the conformational change of NBDs

transmits to the conformational change in the TMDs to open/close the channel is unclear.

Serohijos et al. (2008) constructed a molecular model of CFTR based on its homology to

Sav1866, and they were able to provide some insights into the coupling interface between the

NBDs and the TMDs using this model. They predicted that there are interdomain interactions

between the second intracellular loop (ICL2) in TMD1 and NBD2 and between the fourth

intracellular loop (ICL4) in TMD2 and NBD1. Moreover, they conducted cysteine cross-

linking experiments to investigate the importance of ICL2-NBD2 and ICL4-NBD1 interfaces

to the regulation of channel gating. Single channel activity measurements revealed that

channel activity is restricted upon formation of covalent cross-links between cysteines on

either side of these interfaces, suggesting that both of these interfaces are crucial to the

transmission of regulatory signals. The importance of ICL4 in channel gating was

investigated by Seibert et al. (1996). They reconstructed the eighteen known CF-associated

point mutations in ICL4, and conducted single-channel patch-clamp analysis on the six

mutants that were able to mature. It was found that the mutant channels displayed a

decreased open probability and a Cl- conductance similar to wildtype, suggesting that ICL4

plays an important role in the regulation of channel gating. Also it should be noted that

ΔF508, the most common CF mutation, results in mutant channels with low opening

probability at the cell surface. The F508 residue is predicted to lie at the interface between

NBD1 and ICL4 (Reviewed by Schmidt et al., 2011). Therefore, this can be considered

another evidence supporting the role of the NBD1-ICL4 interface in coupling ATP binding

15

and hydrolysis in the NBDs to channel activity of the TMDs.

1.5 BIOSYNTHESIS AND DEGRADATION OF THE CFTR PROTEIN

The biosynthesis of CFTR starts with the transcription of the CF gene into RNA in

the nucleus. The RNA undergoes splicing, a process in which the noncoding introns are

removed, to produce messenger RNA, which leaves the nucleus, and is translated to an

immature protein in the endoplasmic reticulum (ER) with the help of ribosomes.

CFTR contains two N-linked glycosylation sites in the extracellular loop between TM

segments 7 and 8. Glycosylation is a co-translational event taking place in the ER. When the

glycosylation consensus sequence Asn-X-Ser/Thr, where X is any amino acid except proline,

is at least 12 to 14 residues from the ER membrane, the oligosaccharyltransferase complex

(OST complex) binds to the nascent polypeptide and catalyzes the transfer of a

(Glucose)3(Mannose)9(N-acetylglucosamine)2 group from a dolichol pyrophosphate donor to

the Asn residue (Nilsson and von Heijne, 1993). In the normal folding pathway of wildtype

CFTR, two glucose residues from the oligosaccharide are trimmed by glucosidases I and II,

and the monoglucosylated oligosaccharide structure is recognized by calnexin, a

transmembrane ER chaperone that aids in the protein folding and protects the protein from

aggregation (Reviewed by Amaral, 2005; Reviewed by Farinha and Amaral, 2005).

Folding of the CFTR protein is a complex process involving tertiary folding of

cytosolic domains co-translationally and assembly of TM segments to establish proper

domain-domain contacts post-translationally (Kim et al., 2012). There are molecular

chaperones present in the ER and the cytosol, which ensure proper folding of the protein by

making transient interactions with the nascent polypeptides. Experimental evidence has

16

suggested that many chaperones play important roles in the folding of CFTR. Pulse-chase

and coimmunoprecipitation studies conducted by Yang et al. (1993) revealed transient

association between the cytoplasmic 70kDa heat shock protein (Hsp70) and core-

glycosylated forms of immature CFTR. Hsp70 forms a complex with its co-chaperone

human DnaJ 2 (Hdj-2), and together they facilitate co- and post-translational folding of

CFTR and stabilize NBD1. The interaction of incompletely folded, core-glycosylated CFTR

with the transmembrane ER chaperone calnexin was reported by Pind et al. (1994), who

coimmunoprecipitated pulse-labeled immature CFTR with calnexin from cells transfected

with CFTR. Calnexin interacts with the core-glycosylated forms of immature CFTR until the

glucose from the monoglucosylated oligosaccharide is trimmed by glucosidases II (Reviewed

by Amaral, 2005; Reviewed by Farinha and Amaral, 2005). Mutations such as ΔF508 alter

the interactions of CFTR with the chaperones and cause problems. For instance, Meacham et

al. (1999) have shown that sites necessary for the interaction between NBD1 and R domain

is buried and thus the formation of NBD1-R domain interaction is prevented as a result of

prolonged interaction of ΔF508 CFTR with the Hdj-2/Hsp70 complex. Similarly, prolonged

interaction of calnexin with ΔF508 CFTR leads to ER retention of the mutant protein

(Okiyoneda et al., 2004).

CFTR escapes the ER quality control and is trafficked to the Golgi via COPII-coated

vesicles where it undergoes complex glycosylation if it is folded properly at this stage,

otherwise it is reglucosylated by UDP-glycoprotein glucosyltransferase and retained in the

ER (Reviewed by Amaral, 2005; Reviewed by Farinha and Amaral, 2005). If the protein

remains misfolded after repeated reglucosylation, it is targeted for degradation through the

ER-associated degradation (ERAD) pathway (Reviewed by Molinski et al., 2012). The

17

ubiquitin-proteasome proteolytic pathway is the dominant pathway for degradation of

misfolded CFTR. The misfolded protein is retro-translocated from the ER to the cytosol, and

marked post-translationally by a cytosolic ubiquitin ligase complex containing E3 CHIP.

CHIP recognizes and forms a complex with Hsc70, a chaperone which interacts with the

immature form of CFTR to help the folding process, and remains attached to misfolded

CFTR. Misfolded CFTR is ubiquitinated by the Hsc70 CHIP complex and transported to the

26S proteasome, a multi-protein complex where degradation takes place.

The biosynthesis and maturation of CFTR can be monitored by a difference in

mobility on SDS-PAGE gels. CFTR can exist as three different molecular weight forms on

sodium dodecyl sulphate (SDS) polyacrylamide gel electrophoresis (PAGE) – 120kDa,

170kDa, and 190kDa, corresponding to nonglycosylated CFTR, core-glycosylated CFTR,

and mature CFTR with complex glycosylation, respectively. The presence of a 170kDa band

on an SDS-PAGE gel indicates protein undergoing core-glycosylation in the ER, while the

presence of a 190kDa band indicates protein undergoing complex glycosylation in the Golgi.

In addition, it is also possible to monitor changes in the glycosylation state of CFTR by

enzymatic cleavage with endoglycosidase H and endoglycosidase F. Endoglycosidase H only

cleaves core-glycosylated proteins, while endoglycosidase F is capable of cleaving both core-

glycosylated and complex-glycosylated proteins.

18

1.6 CFTR GENE MUTATIONS AND THEIR CONSEQUENCES AT THE

CELLULAR LEVEL

More than 1900 disease-causing CF gene mutations have been identified to date

(Reviewed by Derichs, 2013). These mutations have been grouped into five classes

according to the primary mechanism underlying the impaired chloride conductance (Welsh

and Smith, 1993) (Table 1). Mutations, such as G542X and R553X (where X is any amino

acid), which result in splice site abnormalities, nonsense mutations or frameshift mutations

leading to premature termination of mRNA translation and ultimately production of a

truncated and mostly non-functional CFTR, are class I mutations. Class II mutations, such as

ΔF508, result in misfolded CFTR protein that is recognized by the cell quality control

mechanism and subsequently degraded instead of getting trafficked from the endoplasmic

reticulum to the Golgi complex and then to the plasma membrane. Class III mutations, such

as G551D, are mutations in the nucleotide-binding domains that affect the direct binding of

intracellular ATP, which result in defective regulation of the channel, and thus, there is no

CFTR function present although full-length CFTR protein is being properly trafficked and

incorporated into the plasma membrane. Class IV mutations, such as R334W, are mutations

in the membrane-spanning domains that affect the channel open probability or the rate of ion

flow, which result in reduced channel conductance. Therefore, despite the proper production,

processing, and regulation of the CFTR protein, there is reduced CFTR function present.

Class V mutations are splice site mutations involving transcription dysregulation, which lead

to slower than normal mRNA splicing and thus decrease the amount of otherwise normal

CFTR protein at the plasma membrane (Reviewed by Kerem, 2005).

19

Table 1 Classes of CFTR Mutations that cause cystic fibrosis.

Class Defect Examples

I splice site abnormalities, nonsense mutations or

frameshift mutations

G542X, R553X

II processing defects resulting in misfolded CFTR

protein

ΔF508

III defective regulation of the channel G551D

IV reduced channel conductance due to mutations

affecting the channel open probability or the rate of

ion flow

R334W

V slower than normal mRNA splicing due to splice site mutations

3120+1G>A (Splice-site mutation in

gene intron 16)

Note: X is any amino acid

20

The most common CF mutation is ΔF508, a class II mutation found in at least one

allele in 90% of CF patients (Bobadilla et al., 2002). Cheng et al. (1990) have conducted

experiments with COS-7 cells to show that cells transfected with vectors containing a ΔF508

cDNA do not express mature, fully glycosylated CFTR. CFTR is an N-linked glycoprotein,

and thus, maturation of the protein can be easily monitored by a shift in size due to the

addition of complex carbohydrate in the Golgi complex when conducting immunoblot

analysis. Based on their results, Cheng et al. (1990) suggested that the protein degradation

machinery detects the misfolded ΔF508 CFTR as having an altered structure compared to the

wildtype, and degrades it. ΔF508 CFTR is degraded either in the ER or in the proteasome

via the ER-associated degradation (ERAD) pathway, instead of getting transported to the

Golgi complex where carbohydrate processing to complex-type glycosylation occurs, and

therefore, only an incompletely glycosylated version of the protein was detected.

The crystal structure of mouse NBD1 was solved by Lewis et al. (2004). It was found

that Phe508 is located at the surface of NBD1 in a region called the α-domain. The side chain

of Phe508 plays an important role in mediating the interaction between NBD1 and TMD2.

The absence of Phe508 leads to an alteration of the length of the α-domain and the

orientation of the residues within it, which ultimately results in improper packing of NBD1

with TMD2 and disrupted association of TMD1 with TMD2 – interaction of NBD1 with

TMD2 is required for the association of TMD1 with TMD2 (Lewis et al., 2004).

Furthermore, cysteine mutagenesis and thiol cross-linking analysis conducted by Chen et al.

(2004) have shown that the deletion of Phe508 abolishes the ability of TMD1 and TMD2 to

be cross-linked to each other. Not only the side chain of Phe508 is necessary for post-

translational formation of intramolecular contacts between the domains of CFTR, the

21

backbone of Phe508 is critical to NBD1 folding efficiency. Thibodeau et al. (2004)

investigated the importance of Phe508 by examining the effects of introducing missense

mutations at this position on the folding of isolated NBD1 in vitro. It was observed that only

the missense mutation F508W affected the folding of the isolated NBD1 – NBD1 folded

poorly at all temperatures when the tryptophan substitution was made. This observation

suggested the important role the peptide backbone of Phe508 plays in proper folding of the

NBD1 domain. Therefore, ΔF508 CFTR is arrested at two different stages – ΔF508 CFTR

with misfolded NBD1 gets degraded rapidly co-translationally in the endoplasmic reticulum,

and ΔF508 CFTR with defective domain-domain contacts is targeted by Hsc70 CHIP E3

ubiquitin ligase for proteasome degradation (Reviewed by Fan et al., 2012).

A minor proportion of the ΔF508 CFTR is able to mature and get trafficked to the

plasma membrane, where it experiences two other problems – defects in gating and a faster

turnover compared to wildtype CFTR (Dalemans et al.1991; Lukacs et al., 1993). Dalemans

et al. (1991) expressed ΔF508 CFTR in Vero cells using recombinant vaccinia virus and

measured channel currents using the whole-cell patch-clamp technique. It was observed that

ΔF508 CFTR exhibited a decreased open probability compared to wildtype CFTR, although

it displayed conductance, anion selectivity and open-time kinetics identical to those of

wildtype CFTR. The relatively shorter residence time of ΔF508 CFTR in the plasma

membrane was suggested by Lukacs et al. (1993), who expressed ΔF508 and wildtype CFTR

in Chinese hamster ovary cells to compare their functional half-lives at the plasma membrane.

The turnover of wildtype and ΔF508 CFTR were assessed by estimating plasma membrane

cAMP-sensitive chloride permeability by membrane potential measurement using bis-oxonol

DiSBAC2 – an anionic voltage-sensitive fluorescent probe that exhibit enhanced fluorescence

22

when cells are depolarized – after blocking protein synthesis with cycloheximide. It was

observed that ΔF508 CFTR has a much higher turnover rate than wildtype CFTR.

The ΔF508 mutation thus seems to cause three major problems – defective folding

and trafficking of CFTR to the cell surface, instability at the cell surface, and impaired

channel activity compared to wild-type CFTR.

1.7 CLINICAL MANIFESTATIONS AND DIAGNOSIS OF CYSTIC FIBROSIS

CF affects multiple organ systems, as CFTR is located at the apical surfaces of

epithelial cells in multiple tissues including the liver, pancreas, intestine, sweat glands, and

lungs, where it plays an important role in determining transepithelial salt transport, fluid flow,

and ion concentrations. In CF patients, normal hydration of the epithelial surfaces is

disrupted due to the lack of chloride channel activity of mutant CFTRs (Gadsby et al., 2006).

Therefore, clinical manifestations of typical CF are chronic obstructive lung disease,

exocrine pancreatic insufficiency leading to malabsorption, sweat gland salt loss, and male

infertility due to absent or altered vas deferens (Reviewed by Zielenski, 2000). Besides these

classical symptoms, other signs of CF can vary from patient to patient, depending on the

severity of the disease. The five classes of CF mutations affect CFTR through different

molecular mechanisms – classes I and V affect CFTR production, and classes II, III, and IV

affect CFTR processing, regulation, and conduction, respectively (Welsh and Smith, 1993).

As a result, the amount of functional CFTR present at the apical membrane varies for the

different CFTR mutations, and the types of CF gene mutations partly determine the severity

of CF symptoms (Reviewed by Zielenski, 2000). It has been found in many studies that

individuals within the same family and carrying the same CF mutation could exhibit different

23

clinical features and clinical course of CF, and thus, it has been suggested that secondary

genetic factors (putative CF modifiers genes), environmental factors, and other non-genetic

factors also likely influence the severity of CF (Reviewed by Zielenski, 2000). Results from

geonotype-phenotype studies assessing the correlation between CF mutations and clinical

outcome characterized by symptoms, severity, and time course conducted by different groups

have shown that CFTR genotype is significantly correlated with exocrine pancreatic function

status of CF patients (Borgo et al., 1990; Kerem et al., 1990; Johansen et al., 1991; Kristidis

et al., 1992; Santis et al., 1990, 1992). Radivojevic et al. (2001) tested whether CFTR

genotype is a good predictor of exocrine pancreatic function by analyzing thirty-two CF

patients with two identified CF gene mutant alleles. They found that thirty-one of the thirty-

two CF patients studied (96.88% of the CF patients studied) were pancreatic insufficient, and

thus, their study supported the hypothesis that exocrine pancreatic function status of CF

patients is genetically determined by their CF gene mutant alleles. Unlike pancreatic function,

the genotype-phenotype correlation for pulmonary function is not as clear. Some studies

reported statistically significant correlations between CFTR genotypes and pulmonary

function (Kerem et al., 1990; Johansen et al., 1991), while others reported otherwise (Santis

et al., 1990; Campbell et al., 1991; Burke et al. 1992; Borgo et al. 1993; Marostica et al.

1998). These discrepancies could be explained by differences in experimental design,

clinical parameters chosen to measure, and the size and demographics of the population

under study. Nevertheless, the severity of pulmonary diseases in CF patients cannot be

reliably determined based on their CF gene mutant alleles. Schechter (2003) conducted a

cross-sectional study to investigate the relationship between the socioeconomic status of a

CF patient and the severity of CF that the patient suffers from. Education attainment, income,

24

and Medicaid insurance status were used as measures of socioeconomic status. Results from

the study have suggested that patients with low socioeconomic status suffered more severe

consequences of CF. The exactly reason is not clear, but other studies have suggested that

worse health and greater mortality is generally associated with the low socioeconomic status

group (Jolly et al., 1991; Mackenbach et al., 1997), and furthermore, healthy Canadian

school children with low socioeconomic status scores display a decreased pulmonary

function compared to their peers with high socioeconomic status scores (Dismissie et al.,

1996). Therefore, it can be concluded that even though CF is a classical Mendelian

autosomal recessive disease, the course of the disease and its clinical presentation could be

influenced by the environment in which the patient lives.

There are three common tests used to diagnose cystic fibrosis: the newborn screening

test, the sweat test, and the genetic test. The newborn screening test is conducted on all

newborns forty-eight to seventy-two hours after birth, and it detects 95% of the newborn

with CF(Genetics in Family Medicine: The Australian Handbook for General Practitioners,

2007). The screening test is based on measurement of immunoreactive trypsinogen in

neonatal bloodspot samples. CFTR common mutation analysis is done on the same bloodspot

sample if elevated immunoreactive trypsinogen is detected, and the presence of two CFTR

gene mutations lead to the diagnosis of CF. Newborns with only one CFTR gene mutation

present is diagnosed to be CF carriers. Since only mutations with high frequencies, such as

ΔF508 CFTR, are screened for, it is possible for newborns with low frequency CF mutation

to be missed by the newborn screening test. Therefore, it is necessary to conduct another test,

the sweat test, to confirm the diagnosis of CF or CF carriers, and to detect those newborns

missed by the screening test. The sweat test, commonly performed at any time from one

25

week of age, is based on measurement of the amount of chloride in sweat. CFTR is located at

the apical surfaces of epithelial cells in the sweat glands, where it plays an crucial role in

determining transepithelial salt transport, fluid flow, and ion concentrations (Reviewed by

Rowe, 2005). The chloride concentration in the sweat of children with CF can be two to five

times higher than the normal chloride concentration in the sweat of healthy children

(Genetics in Family Medicine: The Australian Handbook for General Practitioners, 2007).

The sweat test is convenient and fast, with no needles involved. It can be done in less than an

hour, and results can be obtained on the same day the test is performed. A genetic test could

be further performed to confirm individuals with a positive sweat test result.

1.8 TREATING THE BASIC DEFECT OF CYSTIC FIBROSIS

In vitro studies conducted by Ramalho el al. (2002) on human nasal epithelial cells of

CF patients have shown that achieving as little as 5% of the normal level of wildtype CFTR

activity is sufficient to eliminate the severe pulmonary complication of the disease. Other in

vivo studies have suggested that achieving 10 to 35% of the normal level of wildtype CFTR

activity is necessary (Kerem, 2004). The major disease manifestations can be eliminated with

small amounts of functional CFTR protein at the cell surface, and therefore, the goal of CF

therapy is to promote proper folding of the mutant CFTR, to boost the functional activity of

the CFTR protein trafficked to the plasma membrane, and to stabilize the CFTR protein at

the cell surface.

26

1.8.1 GENE THERAPY

Cystic fibrosis is a monogenic autosomal recessive disease, and thus, theoretically,

gene therapy is the most effective method for correcting the defects. Furthermore, CFTR

gene have been identified, cloned, and characterized by Riordan et al. (1989), the therapeutic

CFTR gene can be easily delivered to the respiratory tract and the lungs, the most affected

organ in CF patients, without any intervention procedures, and low levels of expression of

the normal gene is necessary to correct the cystic fibrosis phenotype. Therefore, it is clear

that gene therapy holds great promise for treating CF. However, an acceptable vector that can

be used to deliver the normal gene to the lungs needs to be first identified. Furthermore, host

specific and non-specific immune responses generated against the foreign therapeutic CFTR

protein is a potential problem that needs to be considered (Reviewed by Proesmans and

Vermeulen, 2008). Figuero et al. (2007) predicted the probability for CFTR to trigger host

cellular immune responses in ΔF508 homozygote patients using the MHC-binding prediction

programs. They have identified a number of potential CD4- and CD8-specific T cell epitopes

within the wildtype CFTR containing the F508 residue, suggesting that there is the

possibility for the injected CFTR to initiate immune responses, and the probability of such

immune responses depends greatly on the activation of T cells specific for the epitopes

within the wildtype CFTR. Immunological mechanisms that might be activated upon

delivery of the vector carrying the therapeutic gene to the respiratory system include the

ingestion of the adenoviral vector by alveolar macrophages (Worgall et al., 1997), and the

initiation of helper T cells dependent humoral immune responses resulting in the generation

of neutralizing antibodies against the vector (Ferrari et al., 2003). Many different approaches

have been developed and utilized to reduce the immunological responses against the vector

27

carrying the therapeutic gene. For instance, in vivo experiments done with mice have shown

that the use of cyclophosphamide, an immunosuppressant drug, effectively prolongs

transgene expression and allows repeated administration of an adenoviral vector (Jooss et al.,

1996). However, the use of immunosuppressant drugs such as cyclophosphamide is

impossible in the clinical setting, as it would reduce the immune responses that protect the

lung cells against foreign particles present in the air to result in accumulation of pathogenic

bacteria in the lungs of CF patients (Kotzamanis et al. 2013). Recent in vivo studies done

with CF mice have suggested that the use of Lentiviral vectors, a member of the Retroviridae

family, is capable of integrating into the host genome and correct the basic

electrophysiological defect, while allowing for long-lasting gene expression without the use

of immunosuppressant drugs (Castellani and Conese, 2010).

An ideal vector system for CF patients not only needs to be able to carry the

therapeutic gene into host cells and ensure it is expressed with an efficiency enough to

correct the CF phenotype, it also should be able to escape the host immune system to allow

long duration of expression and the potential to be safely re-administered. There are two

main types of vector systems currently in clinical trials: viral vectors and cationic liposomes.

Viral vectors, such as Lentiviral vectors, incorporate the CFTR cDNA into the viral genome,

enter host cells, and allow for high levels of gene expression (Castellani and Conese, 2010).

Cationic liposomes are positively charged lipososmes capable of forming a complex with

plasmid DNA encoding CFTR. The cationic liposome-plasmid DNA complexes enter the

host cells and allow for expression of the gene. The levels of CFTR expression using the

cationic liposome-mediated gene transfer method have been relatively poor compared to that

using the viral vector systems, but the cationic liposome-mediated gene transfer method has

28

been found to generate a lower immune response than the viral vector systems (Castellani

and Conese, 2010). Last year, the UK Cystic Fibrosis Gene Therapy Consortium received £3

million in funding from the Medical Research Council and the National Institute for Health

Research funded, and they initiated the largest gene-therapy trial using cationic liposomal

gene delivery systems (Alton et al., 2013). The clinical trials are still on-going, and gene

therapy remains a promising potential treatment for CF patients.

1.8.2 INDIRECT RESCUE APPROACHES

Indirect rescue approaches are methods of promoting proper folding and stabilizing

protein conformation, not by interacting directly with the protein, but rather by alterations in

chaperone interactions, trafficking/recycling pathways, or degradation pathways. Incubating

the cells expressing the mutant CFTR at low temperatures is an example of indirect rescue

approaches. Denning et al. (1992) studied the effect of temperature on the processing of

ΔF508 CFTR and found that the processing defect can be corrected to yield more functional

CFTR in the plasma membrane when the incubation temperature is reduced. Another

example is expressing the mutant protein in the presence of chemical chaperones such as

glycerol. Sato et al. (1996) have shown through in vitro experiments that glycerol can exert

dose- and time-dependent and fully reversible effects on ΔF508 CFTR polypeptides to

stabilize immature core-glycosylated ΔF508 CFTR and thereby increase the processing of

core-glycosylated, endoplasmic reticulum – arrested ΔF508 CFTR into the fully glycosylated

form. Although we are able to obtain partially functional ΔF508 CFTR at the plasma

membrane by treatments such as low temperature protein expression and addition of glycerol

29

to cell culture medium, the rescued ΔF508 CFTR displays four- to six- fold faster metabolic

turnover at the cell surface compared to wildtype CFTR (Sharma et al., 2004). Furthermore,

since these methods are nonspecific in that they may alter the expression or activity of other

proteins, affect other metabolic pathways and cause side effects, they are unlikely to be of

therapeutic benefit.

It is also possible to rescue ΔF508 CFTR by regulating chaperone expression to either

promote the entrance of mutant CFTR into the secretory pathway or inhibit ER- or

proteasome- associated degradation. It has been proposed that CF arises due to defective

interactions between CFTR and the components of the proteostasis network, which includes

the Hsp90 and Hsp40-Hsc/p70 chaperone/co-chaperone ATPase systems responsible for

CFTR folding and degradation, respectively (Balch et al., 2011). Hsp90 is an abundant

chaperone in cells that functions to prevent protein aggregation and assist protein folding.

Loo et al. (1998) have conducted in vitro experiments with CHO and BHK cells expressing

ΔF508 CFTR to show that Hsp90 can facilitate ΔF508 CFTR folding by interacting directly

with its cytoplasmic domains on the ER surface. Disrupting the interaction between Hsp90

and CFTR using the ansamycin drugs was found to block the maturation of the mutant

protein and greatly accelerate its degradation by the proteasome. Wang et al. (2006) have

suggested the interaction of ΔF508 CFTR with Hsp70 and Hsp90 can be altered by

manipulating the ATP loading and ATPase activating co-chaperones governing the ATPase

activities of Hsp70 and Hsp90. They conducted immunoprecipitation to show that a

reduction in the Hsp90 ATPase activator co-chaperone Aha1 in a lung cell line expressing

ΔF508 CFTR (CFBE41o-) by siRNA silencing alters the interactions of ΔF508 CFTR with

Hsp90 to result in stabilization and increased trafficking. The proteasome-associated

30

degradation can also be inhibited to rescue mutant CFTR. The Hsc70 CHIP E3 ubiquitin

ligase targets ΔF508 CFTR with defective domain-domain contacts for proteasome

degradation. Alberti et al. (2004) have identified the co-chaperone HspBP1, a nucleotide

release factor of Hsc70 which interacts with the ATPase domain of Hsc70, as an inhibitor of

the CHIP ubiquitin ligase. Results from immunoprecipitation, immunofluorescence analysis,

and in vitro assays have revealed that HspBP1 can regulate Hsc70-mediated protein quality

control by cooperatively binding to Hsc70 with CHIP. It is suggested that HspBP1 either

shields Hsc70 and the bound mutant CFTR against CHIP-mediated ubiquitylation or prevent

the CHIP ubiquitin ligase from reaching the ubiquitin attachment sites by inducing

conformational changes, and thereby inhibits the CHIP-mediated ubiquitylation of CFTR to

increase trafficking of wildtype and mutant CFTR to the cell surface. The interactions of

CFTR and the components of the proteostasis network could be modulated by proteostasis

regulator which alter the composition and concentration of the proteostasis network to

correct the primary defects in CF disease (Balch et al., 2008; Hutt et al., 2009; Hutt and

Balch, 2010). Cystamine is one such proteostasis regulators identified by Luciani et al.

(2010). They have shown that defective CFTR causes autophagy inhibition and induces

aggresome formation, and cystamine is capable of triggering autophagy pathways to restore

trafficking of ΔF508 CFTR to the cell surface in vitro.

1.8.3 DIRECT RESCUE AND THE USE OF PHARMACOLOGICAL CHAPERONES

Since boosting CFTR activity in CF patients can help to reduce disease severity,

another possible treatment for CF patients is to directly increase channel activity using

potentiators, promote folding of the protein using correctors, or increase stability of the

31

protein at the cell surface using stabilizers. The use of potentiators, correctors, and stabilizers

are likely to be of therapeutic benefit, since this direct approach provides more specific

rescue compared to indirect rescue approaches involving gross changes to protein-protein

interactions and/or protein-solvent interactions. The use of high-throughput screening

technique for identification of potentially active compounds is rapidly growing (Galietta et

al., 2001).

For Class I mutations, which result in splice site abnormalities, nonsense mutations or

frameshift mutations leading to premature termination of mRNA translation, agents that

increase ribosomal ambiguity and decrease its proofreading efficiency can be used to ensure

complete translation of the full-length protein (Reviewed by Proesmans and Vermeulen,

2008). Aminoglycoside antibiotics such as gentamicin are such agents that allow translation

and expression of full-length CFTR protein, as shown in a double-blind, placebo-controlled,

crossover trial conducted by Wilschanski et al. (2003) with cystic fibrosis patients having

premature stop codons. However, the clinical use of gentamicin is limited by its potential

ototoxicity and nephrotoxicity. More recently, another aminoglycoside antibiotic called

amikacin has been identified to provide more effective suppression of the human G542X-

CFTR stop mutation than gentamicin through studies conducted with a transgenic CF mouse

model (Du et al., 2006). Another such agent that induces ribosomal read-through of

premature stop codons is PTC124, a new chemical compound widely studied in healthy

volunteers and in CF patients. Studies conducted by Welch et al. (2007) have suggested

PTC124 has good oral bioavailability, and its phase II studies in patients with nonsense

mutation-mediated cystic fibrosis are currently in progress.

For Class II CF mutations, such as ΔF508, the mutant CFTR is partially functional

32

when trafficked to the plasma membrane (Sampson et al., 2011). Therefore, a possible

treatment for CF patients with the ΔF508 mutant would be to promote maturation and

trafficking of ΔF508 CFTR using pharmacological chaperones. Pharmacological chaperones

are correctors that are specific for CFTR and are predicted to promote maturation by binding

directly to the misfolded protein. A potential advantage of pharmacological chaperones over

indirect rescue approaches like low temperature rescue is that it may interact with the mutant

protein in the endoplasmic reticulum to yield a more stable conformation at the cell surface.

Another potential advantage of pharmacological chaperones is that they may cause fewer

side effects by not altering the expression or activity of other proteins. Finally, another

advantage of specific correctors is that they would not be substrates of drug pumps such as P-

glycoprotein (P-gp), which could reduce the bioavailability of the corrector by pumping it

out of the body (Loo et al., 2012).

1.9 EXPERIMENTAL EVIDENCE IN SUPPORT OF A DIRECT RESCUE

APPROACH

Recent human clinical trials have demonstrated that the potentiator VX-770 can

enhance CFTR channel activity of mutant CFTRs at the cell surface (Accurso et al., 2010).

Furthermore, screening of chemical libraries has identified numerous compounds that act as

correctors to improve ΔF508 CFTR maturation and trafficking to the cell surface (Kalid et al.,

2010). Many of these compounds have been found to exert their effects by directly

interacting with the domains of CFTR. For instance, Sampson et al. (2011) conducted

differential scanning fluorimetry to show that RDR1 directly interacts with NBD1 of CFTR.

However, the efficiency of rescue of ΔF508 CFTR with correctors identified to date is

33

probably too low for therapeutic application. The best corrector identified to date is VX-809

(developed by Vertex Pharmaceuticals) (Kalid et al., 2010). VX-809 has been shown to

increase CFTR function by increasing the trafficking of ΔF508 CFTR that retains some

functional activity at the cell surface in vitro (Clancy et al., 2011). However, a 28-day phase

IIa clinical trial of VX-809 with adult patients who were homozygous for the ΔF508 CFTR

mutation has revealed that after treatment with daily doses of 100-200 mg of VX-809, there

was a statistically significant reduction in sweat chloride values, but there was no statistically

significant improvement in CFTR function in the nasal epithelium as measured by nasal

potential difference. Furthermore, there was no statistically significant change in lung

function, and no maturation of immature ΔF508 CFTR was detected in any of the rectal

biopsy specimens from VX-809 treated subjects (Clancy et al., 2011).

Experiments performed on P-glycoprotein (P-gp), also known as multidrug resistance

protein 1 (MDR1), have also provided evidence supporting the possibility of using direct

rescue approaches to correct CFTR defects. The P-gp drug pump is another member of the

ABC family of proteins. It is a useful model system for studying defective folding and

trafficking of CFTR processing mutants, as modeling and electron crystallography studies

suggest that P-gp is structurally similar to CFTR (Loo et al., 2007). P-CFTR shares 30%

sequence homology with P-gp (Lallemand et al., 1997). P-gp also contains two NBDs and

two TMDs, but lacks the R domain (Figure 2). The R domain may not be essential for

folding as deletion of residues 708-830 from the R domain of CFTR does not affect protein

maturation (Vankeerberghen et al., 1999). It has been found that the deletion of Tyr490 from

P-gp, which is equivalent to the deletion of Phe508 from CFTR, also inhibits maturation of

the protein (Loo and Clarke, 1997). To be specific, the deletion of Tyr490 from P-gp disturbs

34

the interaction between the first nucleotide binding domain, where the residue Tyr490 is

located, and the first cytoplasmic loop, and thereby results in disrupted packing of the TM

segments (Loo et al., 2002). Furthermore, it was shown that expressing ΔY490 P-gp in the

presence of drug substrates, which bind directly to the transmembrane domains of ΔY490, P-

gp could promote maturation to yield a functional protein at the cell surface. An even more

remarkable finding was that expression of P-gp processing mutants containing mutations in

any domain could be rescued when expressed in the presence of drug substrates (Loo et al.,

1997). Since P-gp and CFTR are structurally similar, we hypothesize that CFTR containing

processing mutations like ΔF508 can be repaired by a P-gp drug-rescue mechanism. The

mechanism of P-gp drug rescue is that drugs specifically bind to the transmembrane domains

of processing mutants to repair defects in packing of the transmembrane segments and

promote domain-domain interactions (Loo et al., 2009).

Experiments previously done in our lab have shown that arginine suppressor

mutations introduced in the TM segments of P-gp can mimic the drug rescue effects to

promote folding of P-gp processing mutants, such as ΔY490 P-gp (Loo et al., 2007). A

suppressor mutation is a second mutation that can counter the phenotypic effects of an

already existing mutation. Arginine is a unique amino acid in that it has a positively charged

side chain, and it is capable of forming up to three hydrogen bonds. It was found that

arginine suppressor mutations introduced into the TM segments of P-gp processing mutants

promoted interdomain or intradomain hydrogen bond interactions between adjacent TM

segments, and thereby, mimicked drug-rescue to promote maturation of P-gp processing

mutants (Loo et al., 2007).

35

Figure 2 Models of CFTR and P-glycoprotein. The 12 transmembrane segments of full-length CFTR or P-glycoprotein are shown as numbered cylinders, and the glycosylation sites

are shown as branched lines. TMD, NBD, and R represent the transmembrane domains, the

nucleotide-binding domains, and the regulatory domain of CFTR, respectively. The locations

of ΔF508 and ΔY490 are indicated. The glycosylation sites are located in the first

extracellular loop of TMD2 in CFTR and the first extracellular loop of TMD1 in P-

glycoprotein. Both F580 of CFTR and Y490 of P-glycoprotein are located in NBD1. The

position of residue Y490 of P-glycoprotein is equivalent to the position of F508 in CFTR,

and ΔY490 in P-glycoprotein is equivalent to ΔF508 in CFTR.

36

It has been found that the introduction of V510D in NBD1 of ΔF508 CFTR partially

corrects the folding defects to promote maturation and stability at the cell surface (Loo et al.,

2010). Other suppressor mutations, such as I539T, G550E, R553Q, and R555K, identified in

NBD1 of ΔF508 CFTR were found to have similar effects as V510D (Reviewed by Schmidt

et al., 2011). Furthermore, introduction of the suppressor mutation I539T into ΔF508 NBD1

was found to completely restore NBD1 conformation and stability (Hoelen et al., 2010). The

identification of suppressor mutations in CFTR suggests the possibility of restoring proper

assembly of ΔF508 CFTR through specific rescue.

1.10 OBJECTIVES

1.10.1 ARGININE SCANNING MUTAGENESIS OF THE TRANSMEMBRANE

SEGMENTS OF CFTR

In this thesis, arginine scanning mutagenesis of the TM segments of CFTR was

performed. Since modeling studies and crystallization studies have suggested that Phe508

from NBD1 is situated next to intracellular loop 4 in TMD2 (Lewis, 2004; Serohijos, 2008),

and thus, the ΔF508 mutation likely disrupts NBD1-TMD2 interactions and thereby disrupts

packing of the TM segments, the TM segments are predicted to be good target sites for

correctors. Knowledge regarding the structure of the TMDs of CFTR will be useful in

developing better correctors and understanding their mechanisms. Furthermore, as mentioned

above, CFTR‟s sister protein, the P-gp drug pump containing the equivalent mutation

(ΔY490), could be repaired by a drug-rescue approach (Loo et al., 1997). The drug substrates

did not rescue CFTR processing mutants, suggesting their specificity against P-gp (Loo et al.,

1997). Moreover, the mechanism of drug-rescue involved direct binding to the

37

transmembrane domains (TMDs) since over 38 arginine suppressor mutations were identified

in TM segments of P-gp that mimicked drug-rescue to promote maturation of processing

mutants (Loo et al., 2007). We hypothesized that CFTR folding defects could be

corrected by introducing arginines in the transmembrane domains of the protein, and

furthermore, CFTR containing processing mutations like ΔF508 can be repaired by a

P-gp drug-rescue mechanism – a mechanism in which drugs specifically bind to the

transmembrane domains of processing mutants to repair defects in packing of the

transmembrane segments and promote domain-domain interactions. To address the

question of whether CFTR processing mutants could be specifically and directly repaired by

a similar drug-rescue approach, arginine mutagenesis was performed on „unstable‟ TM

segments 6, 8, and 12 of CFTR to test for suppressors. These TM segments were chosen

because a study has shown that TM8 and TM12 are the only TM segments that do not insert

well into the ER membrane by themselves, and TM6 requires its natural C-terminal flanking

region for efficient insertion into the membrane (Enquist, 2009). Furthermore, these three

TM segments are among the least hydrophobic in the protein, as judged by the predicted ΔG

values. A study performed by Tector, M. and Hartl, F.U. (1999) has also demonstrated that

TM6 of CFTR is extremely unstable in the lipid bilayer upon membrane insertion. TM6 fails

to act as an efficient anchor sequence in the ER. It is the ribosome-ER translocation

machinery and the cytosolic domains of CFTR that co-operate to inhibit the slipping of TM6

into the ER lumen.

Our objectives in performing arginine scanning mutagenesis of the TM segments of

CFTR were twofold:

(1) To predict the relative positions of the residues in the TMDs of CFTR.

38

(2) To identify arginine suppressor mutations in the TM segments of CFTR.

1.10.2 DIRECT RESCUE OF CFTR PROCESSING MUTANTS USING

CORRECTORS

The use of potentiators, correctors, and stabilizers is likely to be of therapeutic benefit,

since this direct approach provides more specific rescue compared to indirect rescue

approaches involving gross changes to protein-protein interactions and/or protein-solvent

interactions. We hypothesized that pharmacological chaperones confer specificity to

CFTR by binding directly to the protein to promote maturation and enhance stability

of the protein. Since recent human clinical trials have demonstrated that the potentiator VX-

770 can enhance CFTR channel activity of mutant CFTRs at the cell surface (Accurso, 2010),

this thesis focused on identifying approaches to improve the maturation, trafficking, and cell

surface stability of ΔF508 CFTR. As mentioned above, the mechanism of drug-rescue of

ΔY490 P-gp involved specific and direct binding to the TMDs of the protein (Loo et al.,

1997). To investigate the possibility of repairing CFTR processing mutants specifically and

directly by a similar drug-rescue approach, we tested different correctors for their specificity

(identified correctors that rescue CFTR but not P-gp processing mutants) and used truncation

mutants to map the VX-809 rescue site. VX-809 was chosen because it is the best corrector

identified to date for CFTR. It is known that many of the correctors discovered to date for

CFTR exert their effects by directly interacting with the domains of CFTR. For instance,

differential scanning fluorimetry conducted have shown that RDR1 directly interacts with

NBD1 of CFTR (Sampson et al., 2011). Other previous studies have shown that many

correctors, such as VX-325, are non-specific and could rescue P-gp processing mutants

39

(Kalid et al., 2010). Therefore, this thesis identified correctors that do not rescue P-gp

processing mutants and tested if they promote maturation of ΔF508 CFTR into a more stable

protein compared to low temperature rescue.

The objectives of this portion of the research were:

(1) To test whether CF processing mutations, such as H1085R and V232D, reduce

stability

of CFTR.

(2) To identify correctors that specifically rescue ΔF508 CFTR.

(3) To identify the potential interaction sites for corrector molecules by examining the

effect of correctors on the stability of CFTR domains expressed as separate

polypeptides.

40

2 METHODS

2.1 CONSTRUCTION OF MUTANTS

All mutants used in this thesis were constructed by Dr. Tip Loo. Arginine mutations

were introduced into wildtype CFTR cDNA by the method of Kunkel (1985). Plasmids

containing the wildtype CFTR cDNA were transformed into an ung ̄dut ̄strain of E. coli

bacteria (an E. coli strain incapable of breaking down dUTP and removing uracil from newly

synthesized DNA due to deficiency in dUTPase and uracil deglycosidase) to produce single-

stranded DNA with uracil incorporated in place of thymine. This single-stranded DNA was

extracted and incubated with an oligonucleotide containing the desired mutation to generate

double-stranded plasmid consisting one parental non-mutated strand containing uracils and a

mutated strand containing thymines through polymerization reaction cycles. Finally, the

double-stranded plasmid generated was transformed into an E. coli strain carrying the

wildtype dut and ung genes. dUTPase breaks down dUTP in the cells and uracil

deglycosidase removes any incorporated uracil in the plasmid, and thus, nearly all of the

resulting plasmids contain the newly mutated sequence (Kunkel, 1985). Using the same

method, arginine mutations were introduced into H1085R, V232D, or ΔF508 CFTR to create

double mutants. Plasmids expressing half molecules or truncation mutants of CFTR were

constructed using methods described by Chan et al. (2000). The CFTR cDNA coding for

wildtype, H1085R, V232D, and ΔF508 CFTR was inserted into the pcDNA3 expression

vector. The CFTR cDNAs coding for ΔNBD2 CFTR (residues 1-1196), the NH2-terminal

half-molecule (N-half CFTR) (residues 1-633), COOH-terminal half-molecule (C-half CFTR)

(residues 837-1480), TMD1 CFTR (residues 1-388), TMD2 CFTR (residues 837-1196),

41

TMD1+2 (TMD1+2 CFTR) (residues 1-388 plus 837-1196), NBD1 CFTR (residues 387-

646), and the P-glycoprotein cDNA coding for G268V P-gp or G251V Pgp were inserted

into the pMT21 expression vector. Wildtype and ΔF508 CFTR, and CFTR half-molecules

and truncation mutants were modified to contain the epitope tag for monoclonal antibody

A52 at the COOH-terminal end of the protein for easy detection of transfected CFTR rather

than endogenous CFTR. The integrity of the mutated cDNAs was confirmed by DNA

sequencing.

2.2 CELL CULTURE

All expression assays and transfection were conducted on HEK-293 (human

embryonic kidney) cells transiently transfected with the wild-type or mutant cDNAs. BHK

(baby hamster kidney) cells stably expressing the wild-type or mutant protein were used for

cell surface labeling experiments and iodide efflux assays.

HEK-293 cells were grown in Dulbecco‟s modified Eagle‟s media (DMEM) fortified

with 0.1 mM minimum Eagle‟s medium non-essential amino acids, 2 mM L-glutamine, 100

units of penicillin/ml, 100 µg streptomycin/ml, 10% v/v calf serum in 5% CO2 at 37C. Cells

were split to 50% confluency and grown overnight. The following day, cells were transfected

with cDNA coding for the wild-type or mutant CFTR. To transfect one well of a 6-well plate,

the amount of DNA needed to obtain a final concentration of 1 µg/mL was added to 67.5 µL

of H2O. 2.5 M CaCl2 was added to a final concentration of 12.5 mM and mixed by swirling.

A volume of 75 µL of 2X N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid (BES) (50

mM BES, 280 mM NaCl, 1.5 mM Na2HPO4, pH 6.96 with NaOH) was added dropwise. The

42

mixture was allowed to sit for 10 minutes at room temperature, after which 1.5 mL of cell

culture medium was added. For mutant CFTR that show low expression, 1 mM sodium

butyrate was added to the cell culture medium to boost expression. The old medium was

removed from the cells and the medium containing the calcium phosphate-precipitated DNA

was gently added to the well. Cells were then incubated for about 5 hours at 37C, after

which the medium was changed to either fresh medium or medium containing a corrector of

interest and incubated overnight at 37C. The next day, cells were harvested.

BHK cells were grown and transfected the same way as HEK-293 cells, except that

10 cm plates were used instead of 6-well plates, and selection vector, pwl-neo, was added to

transfection media with the DNA of interest at a ratio of 1:20. The next day after transfection,

selection media containing 1 mg/ml active concentration of G418 was applied, and the cells

were incubated at 37C for 10~14 days until colonies started to form. Twenty-four colonies

were picked for each construct. The colonies were allowed to grow in 24-well plates for 3~4

days, after which duplicate colonies were made and were allowed to grow for a couple of

days until confluent. One set of the duplicates was used to run a Western blot (see

WESTERN BLOTTING section), while the other set was used to maintain any positive

colonies. Three colonies that were expressing well were selected and transferred to T75

flasks for each construct. To freeze the cell lines for future use, Nunc cryotubes were used

for storage and 10% dimethyl sulfoxide (DMSO) in DMEM was used as freezing media. The

cells were washed with 5 ml phosphate buffered saline (PBS), and 2.5 mL of 0.25% trypsin

was added to release the cells from the flask. After about 1 min, 8 ml of fresh DMEM was

added. All cells plus media was transferred into a 15 mL conical tube, and spun in a bench

43

top IEC clinical centrifuge at setting #3 (about 4000 rpm) for 3 minutes. The pellet obtained

was suspended in 3 mL of freezing media, and then 1.5 mL was transferred to a Nunc

cryotube. The Nunc cryotubes were placed at -70C for 24 hours, and then transferred to

liquid nitrogen storage.

2.3 CELL SURFACE LABELING

Confluent BHK cells stably expressing the wild-type or mutant protein were washed

four times with phosphate buffered saline (pH 7.4) containing 0.1 mM CaCl2 and 1 mM

MgCl2 (PBSCM), and then treated in the dark with PBSCM buffer containing 10 mM sodium

periodate for 30 minutes at 4C. The cells were then washed four times with PBSCM buffer

and treated with sodium acetate buffer (100 mM sodium acetate buffer, pH 5.5, 1 mM MgCl2

and 0.1 mM CaCl2) containing 2mM biotin-LC-hydrazide for 30 minutes at 20C. The cells

were then washed twice with sodium acetate buffer and solubilized with

tris(hydroxymethyl)-aminoethane (Tris)-buffered saline (100 mM Tris-HCl, pH 7.4 and 150

mM NaCl) containing 1% (w/v) octyl phenoxy polyethoxyethanol (Triton X-100), 0.5% (w/v)

sodium deoxycholate, and 1mM ethylenediaminetetraacetic acid (EDTA). After being placed

on ice for 5 minutes, the cells were transferred to a 1.5 mL tube and were spun at 15,000 rpm

for 5 minutes. The supernatant was collected, and CFTR was immunoprecipitated with 1.1

mg/mL monoclonal antibody A52, subjected to SDS-PAGE on 6.5% gels and biotinylated

CFTR was detected with streptavidin-conjugated horseradish peroxidase and the ChemiDoc

XRS+ imaging system, which is a chemiluminescent detection system by Bio-Rad

44

Laboratories, Inc.

2.4 CYCLOHEXIMIDE CHASE ASSAY

To test if a corrector promoted stability of a CFTR mutant, transiently transfected

HEK-293 cells were grown and transfected as described above (see CELL CULTURE

section). Transfected cells were incubated overnight at 30C in the presence or absence of

corrector after the change of medium. The next day, 0.5 mg/mL cycloheximide was added to

stop protein synthesis and the cells were placed at 37C. Cells were harvested 0, 1, 2, 4, 6, 8,

16, and 24 hours after addition of cycloheximide. 10% DMSO was added and the cells were

frozen to stop protein degradation.

2.5 WESTERN BLOTTING

The expression of wild-type and mutant CFTRs was detected by immunoblot analysis.

Whole HEK-293 and BHK cells transfected with wild-type or mutant cDNAs were

solubilized in 120µL of SDS-PAGE sample buffer containing 50 mM EDTA and 2% β-

mercaptoethanol, and resolved by SDS-PAGE on 6.5%, 10% or 12% gels (15µL of the

samples were loaded into each well). The proteins were transferred to a nitrocellulose

membrane by electroblotting for 50 minutes at 490 milliamps. The nitrocellulose was

blocked in 1% w/v milk powder dissolved in Tris-buffered saline (TBS) (10 mM Tris HCl,

150 mM NaCl, pH 7.5) containing 0.5% (v/v) Tween-20 (TBST) for 15 minutes.

For wildtype and mutant cDNAs modified to contain the epitope tag for monoclonal

antibody A52 at the COOH-terminal end of the protein, the blot was incubated in 1% milk in

45

TBST with serum containing a mouse monoclonal antibody against the A52 tag (1:200

dilution) at 4C overnight. The blots were then washed three times for 5 minutes with TBST

and then incubated in 1% milk in TBST with serum containing an anti-mouse, horse radish

peroxidase-conjugated antibody (1:5,000 dilution) at 4C overnight. After 3 washes of 5

minutes with TBST, the ECL, a chemiluminescent substrate for the horseradish peroxidase

enzyme, was applied to the blots, and CFTR was detected using the ChemiDoc XRS+

imaging system, which is a chemiluminescent detection system by Bio-Rad Laboratories, Inc.

For wildtype and mutant cDNAs that do not contain an A52-epitope tag, the blot was

incubated in 1% milk in TBST with serum containing a rabbit polyclonal antibody against

CFTR (1:5,000 dilution) at 4C overnight. The blots were then washed three times for 5

minutes with TBST and then incubated in 1% milk in TBST with serum containing an anti-

rabbit, horse radish peroxidase-conjugated antibody (1:20,000 dilution) at 4C overnight.

After 3 washes of 5 minutes with TBST, the ECL, a chemiluminescent substrate for the

horseradish peroxidase enzyme, was applied to the blots, and CFTR was detected by

chemiluminescence using the ChemiDoc XRS+ imaging system.

To scan and quantitate the gel lanes, the Image Lab image acquisition and analysis

software from Bio-Rad Laboratories, Inc. and a Windows computer were used.

46

2.6 IODIDE EFFLUX ASSAY

Stably transfected BHK cells that were grown and transfected as described above (see

CELL CULTURE section) were used for the iodide efflux assay. The culture medium was

aspirated from the 80~90% confluent cell monolayer, and the cells were gently washed three

times with 2 mL of an iodide loading buffer (136 mM sodium iodide, 4 mM potassium

nitrate, 2 mM calcium nitrate, 11 mM glucose and 20 mM HEPES, pH 7.4 with NaOH)

warmed to 37C. The cells were incubated in 2 mL of the loading buffer for one hour in the

dark at room temperature. Following the incubation period, the loading buffer was removed

by slowly aspirating and the cells were gently washed ten times (1 minute each) with 2 mL

of an iodide free efflux buffer (136 mM sodium nitrate, 4 mM potassium nitrate, 2 mM

calcium nitrate, 11 mM glucose and 20 mM HEPES, pH 7.4) warmed to 37C. The cells

were equilibrated in 1 mL of iodide free efflux buffer for one minute at room temperature,

after which the buffer was removed and replaced with 1 mL of fresh iodide free buffer. The

removed samples of efflux buffer were collected in 24-well plates, and measurements were

done using an iodide sensitive electrode to establish a stable baseline. After three rounds of

efflux buffer collection, a stimulating buffer (4 mM potassium nitrate, 2 mM calcium nitrate,

11 mM glucose and 20 mM HEPES, pH 7.4) containing 200 µM IBMX, 10 µM forskolin, 50

mM genistein, and 200 µM cpt-cAMP was added at one minute intervals for 12 minutes. The

removed samples of stimulating buffer were collected in 24-well plates, and measurements

were done using an iodide sensitive electrode. An iodide concentration versus voltage

standard curve was constructed by measuring the electrode value (in mV) in solutions

containing from 10 mM to 1 µM I-, and the equation of this line was then used to determine

47

the amount of iodide in samples of efflux buffer from individual experiments. Iodide

concentration versus time was plotted to generate a time-course of iodide efflux from BHK

cells expressing wild-type or mutant CFTRs.

48

3 RESULTS

3.1 ARGININE MUTAGENESIS OF CFTR TM SEGMENTS

Currently there is not a high-resolution structure of the full-length human CFTR protein.

However, modeling studies and crystallization studies have predicted that Phe508 from NBD1 is

situated next to the fourth intracellular loop (ICL4) in TMD2 (Lewis et al., 2004; Serohijos et al.,

2008). The most common CF mutation, ΔF508, likely disrupts packing of the transmembrane

segments by disrupting NBD1-TMD2 interactions (Chen et al., 2004). Therefore, the

transmembrane segments are predicted to be good target sites for correctors, and knowledge

regarding the structure of the TMDs of CFTR will be useful in developing better correctors and

understanding their mechanisms. Arginine is a unique amino acid in that it remains charged in

nonpolar environments, and its side chain is capable of forming up to three hydrogen bonds (Li

et al., 2008). Previous work on P-glycoprotein (P-gp), another member of the ABC transporter

family that is structurally similar to CFTR, has shown that arginine suppressor mutations

introduced into the TM segments of P-gp processing mutants, such as ΔY490 P-gp, which is

equivalent to ΔF508 CFTR, mimicked drug-rescue to enhance maturation by promoting

interdomain or intradomain hydrogen bond interactions between adjacent TM segments (Loo et

al., 2007). We hypothesized that CFTR containing processing mutations like ΔF508 can be

repaired by a P-gp drug-rescue mechanism – a mechanism in which drugs specifically bind to the

transmembrane domains of processing mutants to repair defects in packing of the transmembrane

segments and promote domain-domain interactions. If CFTR processing mutants can be repaired

by a similar drug-rescue mechanism, then we predict that some arginines introduced into the TM

segments will act as suppressors to promote maturation of the mutant protein.

49

3.1.1 MAPPING THE STRUCTURE OF CFTR TMDs AND TESTING WHETHER

ARGININES INTRODUCED IN THE TMDs OF WT-CFTR PROMOTE MATURATION

Arginine-scanning mutagenesis of TM6, TM8, and TM12 of CFTR was performed.

These TM segments were chosen because a study has shown that TM8 and TM12 are the only

TM segments that do not insert well into the ER membrane by themselves, and TM6 requires its

natural C-terminal flanking region for efficient insertion into the membrane (Enquist, 2009). It

has been suggested that TM6 of CFTR is extremely unstable in the lipid bilayer upon membrane

insertion. It fails to act as an efficient anchor sequence in the ER, and it is the ribosome-ER

translocation machinery and the cytosolic domains of CFTR that co-operate to prevent it from

slipping into the ER lumen (Tector, 1999). Furthermore, cysteine mutagenesis and thiol cross-

linking analysis conducted by Chen et al. (2004) have shown that the ΔF508 mutation abolishes

the ability of TM6 and TM12 to be cross-linked to each other. Therefore, we predicted that some

arginines introduced into TM6, TM8, and TM12 of CFTR would act as suppressor mutations to

stabilize and promote the maturation of CFTR processing mutants, such as ΔF508 CFTR, by

forming interdomain or intradomain hydrogen bond interactions between adjacent TM segments.

The first step was to perform arginine mutagenesis of wildtype CFTR to identify locations

where arginines would not inhibit maturation and test models of CFTR structure. Arginines that

did not inhibit maturation would then be introduced into CFTR processing mutants to test if they

act as suppressors.

To perform arginine scanning mutagenesis of TM6, TM8, and TM12, the cDNA of

wildtype CFTR was modified to create a set of mutants that contained one arginine at positions

332–351, 912–927, and 1134–1145. HEK-293 cells were transiently transfected with plasmids

encoding mutant CFTR, and cells were grown overnight at 37C to allow for expression of the

50

protein. Whole cell extracts of mutant CFTRs were subjected to immunoblot analysis using 6.5%

(w/v) acrylamide gels and polyclonal anti-CFTR antibody (see Methods for details). Figure 3A

shows the immunoblot results for the mutants. The glycosylation of CFTR, monitored by a

difference in mobility of SDS-PAGE gels, served as an indicator of the maturation state of CFTR.

The presence of a 170 kDa band on a SDS-PAGE gel indicated immature protein that was core-

glycosylation in the ER, while the presence of a 190 kDa band indicated mature protein that have

been complex-glycosylated in the Golgi. The conversion of the immature 170 kDa protein to the

mature 190 kDa protein is termed “maturation”. The ratio of mature CFTR to total CFTR was

determined for each mutant CFTR and the wildtype CFTR and was used as a measure of steady-

state maturation efficiency (Figure 3B). Figure 3C shows the positions of the residues in the TM

segments as α-helical wheels and the effect of arginine mutations at various positions on

maturation of CFTR. The most common effect of introducing arginines into TM6, TM8, and

TM12 of wildtype CFTR was to reduce the level of mature protein.

Arginine residues introduced in the TMDs of CFTR were observed to have different

effects on the maturation of the protein. They were seen to completely inhibit maturation and/or

decrease yield of CFTR protein (190 kDa undetectable in cells), partially inhibit maturation (both

170 and 190 kDa detectable in cells, with a decreased relative level of 190 kDa CFTR), or have

little or no effect on maturation. For instance, immunoblot results (Figure 3A) show that S341R

partially inhibited maturation (i.e. the S341R CFTR mutant showed lower steady-state

maturation efficiency than wildtype CFTR), V920R completely inhibited maturation (i.e. the

V920R CFTR mutant had a steady-state maturation efficiency close to zero), and M348R only

had a small effect on maturation (i.e. the M348R CFTR mutant showed the 190kDa protein as

the major product). The V920R mutation may have inhibited maturation because it is predicted

51

Figure 3 Effect of arginine mutations on maturation of CFTR. Wildtype CFTR or mutant

CFTRs containing arginines at various positions in predicted TM segments 6, 8, or 12 were

expressed in HEK cells, and whole cell SDS extracts of cells were subjected to immunoblot

analysis (A). The positions of the mature CFTR (190 kDa), and immature CFTR (170 kDa) are

indicated. The amount of mature CFTR relative to total CFTR (% mature) was quantitated for

each mutant CFTR and was used as a measure of steady-state maturation efficiency (B). Each

value is the mean ± SE. (n=4). The positions of the residues in the TM segments as α-helical

wheels and the effect of arginine mutations at various positions on maturation of CFTR are

shown (C). Arginine mutations that inhibit or have a neutral effect on maturation are shown as

white circles or gray circles, respectively. (D) Schematic models of CFTR. TM6, 8, and 12 are

shown in white, light grey, and dark grey, respectively. Amino acid residues which are potential

suppressor mutations are indicated. The structure was generated and viewed using the PyMOL

Molecular Graphics System (DeLano, 2002).

52

53

54

55

to lie on the lipid face of TM8. By contrast, M348 is predicted to face the aqueous pore of CFTR,

and thus, mutating it to an arginine may have less effect on protein folding and maturation of the

protein. The S341R mutation may have partially inhibited maturation because it is predicted to

reside at the TM3-TM6 boundary.

It was observed that some arginine mutants, such as C343R CFTR, yielded very low

levels of mature and immature protein (i.e. both 170 and 190 kDa CFTR protein bands hardly

detectable in cells), while others, such as Y913R CFTR, yielded very high levels of mature and

immature protein. Furthermore, there were some arginine mutants, such as T338R and T925R

CFTRs, which displayed high levels of immature protein compared to mature protein. A possible

explanation for these observations is that the introduction of an arginine mutation into the TM

segment of CFTR can have different effects on protein folding and stability. Mutations such as

C343R possibly cause severe folding defects, and as a result, the misfolded protein either gets

rapidly degraded co-translationally in the ER or is targeted for proteasome degradation.

Therefore, very low levels of mature and immature protein were observed. Mutations such as

T338R and T925R possibly affect the stability of the mature protein at the cell surface. These

arginine mutants undergo core-glycosylation in the ER as normal, but the mature protein

generated after the mutants undergo complex glycosylation in the Golgi and get trafficked to the

cell surface is unstable and turned over rapidly. As a result, high levels of immature protein

compared to mature protein were observed.

It was observed that I344R, M348R, S912R, Y913R, Y914R, V915R, F916R, and

Q1144R CFTRs showed maturation efficiencies comparable to wildtype CFTR. The positions of

these residues are highlighted in Figure 3D, which shows a schematic model of CFTR generated

and viewed using the PyMOL Molecular Graphics System (DeLano, 2002). These arginine

56

mutations may have had no effect on protein folding and maturation of the protein because they

are pore-lining residues.

3.1.2 IODIDE EFFLUX ASSAYS ON ARGININE MUTANTS

CFTR mutants I344R, M348R, S912R, Y913R, Y914R, V915R, F916R, and Q1144R

were observed to yield the mature 190kDa form of CFTR as the major product (Figure 3). These

arginine mutants are candidates to act as suppressors that could be introduced into a processing

mutant like ΔF508-CFTR to test if they would act as suppressor mutants. Introduction of such a

relatively large charged arginine side chain at positions in the pore-lining TM segments however,

could inhibit channel activity, and thus, iodide efflux assays were performed to examine channel

function of the arginine mutants.

There are different methods of measuring CFTR channel activity. The whole-cell

arrangement of the patch clamp technique and iodide efflux assay are two commonly used

methods. In this thesis, we chose to use the iodide efflux assay to assess channel function since it

is a relatively fast and convenient method that gives reliable results. Although chloride ion is the

natural substrate of CFTR, CFTR is also capable of transporting iodide upon cAMP stimulation.

Iodide has the advantage over chloride in that very few channels are capable of conducting

iodide current besides CFTR. There are many other chloride transporting proteins on the cell

surface, and therefore, the potential for interfering in CFTR assays is greater and the results are

less reliable with a chloride based method (Rich et al., 1991). A disadvantage of the iodide efflux

assay is that CFTR transports iodide with less efficiency than chloride due to the larger size of

the iodide ion (Tabcharani et al., 1992). Nevertheless, the iodide efflux assay is an easy to use

and reliable method for studying CFTR channel activity, and it was the method used in this

57

thesis.

Measurement of IBMX, forskolin, genistein, and cpt-cAMP stimulated iodide efflux was

performed on BHK cells expressing wildtype or mutant CFTRs. Adherent BHK stable cell lines

were generated and used for the iodide efflux assays, as the assays required many washing steps

of cells attached to the plates. To load the cells with iodide, iodide loading buffer containing

sodium iodide was used to wash and incubate the moderately confluent BHK cells. The cells

were then washed 13 times at 1 minute intervals with iodide efflux buffer containing no sodium

iodide. The last three washes were collected and measurements were done using an iodide

sensitive electrode to establish a stable pre-stimulation baseline. Stimulation buffer containing

IBMX, forskolin, genistein, and cpt-cAMP was then added to the cells to stimulate iodide release

from the cells. Stimulation buffer was collected for twelve cycles to obtain a full activation curve

(see Methods). For each experiment, a control of untransfected cells, which gave no efflux at all,

and wildtype CFTR expressing cells were tested. It was found that cells expressing wildtype,

I344R, S912R, Y913R, Y914R, or Q1144R CFTRs exhibited iodide efflux upon addition of the

stimulation buffer, while untransfected cells and cells expressing M348R, V915R, or F916R

CFTRs gave no efflux at all (Figure 4). To confirm that mutants that gave no efflux was due to

non-functional channels rather than failure to load the cells with iodide, the detergent SDS was

added to untransfected cells and cells expressing mutants that gave no efflux to cause lysis and

release of trapped iodide from cells at the end of the experiments (data not shown).

58

Figure 4 Iodide efflux activity of TM6 (A, B), TM8 (C), and TM12 (D) CFTR mutants.

Iodide efflux assays were performed on BHK cells stably expressing wildtype, mutant CFTR, or

no CFTR (untransfected control). Iodide loading buffer containing sodium iodide was used to

wash and incubate moderately confluent BHK cells. The cells were washed 13 times at 1 minute

intervals with iodide efflux buffer, and the last three washes were collected to establish a stable

pre-stimulation baseline (time -3 to 0) At time 0, stimulation buffer containing IBMX, forskolin,

genistein, and cpt-cAMP was added to start stimulation of the iodide-loaded cells.

59

3.1.3 IDENTIFYING SUPPRESSOR MUTATIONS IN THE TMDs OF CFTR

CFTR mutants I344R, S912R, Y913R, Y914R, and Q1144R were found to exhibit

maturation efficiencies and channel activities comparable to wildtype CFTR, thus we introduced

these mutations into ΔF508 CFTR, V232D CFTR, and H1085R CFTR to test for potential

suppressor mutations that could rescue processing mutants. ΔF508 CFTR, V232D CFTR, and

H1085R CFTR were chosen to test for suppressor mutations because these are processing

mutations located in different domains of CFTR – V232D in TMD1, H1085R in TMD2, and

ΔF508 in NBD1 (Figure 5). Furthermore, these three CFTR processing mutants all yield active

proteins after rescue with correctors. The folding defects of V232D, H1085R, and ΔF508 can be

repaired by the chemical chaperones corr-4a (Caldwell et al., 2011), CFcor-325 (Loo et al.,

2006), and VX-809 (Loo et al., 2012), respectively.

To identify suppressor mutations, mutations (I344R, S912R, Y913R, Y914R, or Q1144R)

were introduced into ΔF508 CFTR, V232D CFTR, or H1085R CFTR cDNAs by site-directed

mutagenesis and the cDNAs were transiently expressed in HEK-293 cells. HEK-293 cells were

transfected with the cDNAs, and the medium was changed five hours later to fresh medium. The

next day, cells were harvested after the change in medium, and whole cell extracts were

subjected to immunoblot analysis using 6.5% (w/v) acrylamide gels and a CFTR polyclonal

antibody (see Methods). Once again, the glycosylation state of CFTR, monitored by a difference

in mobility of SDS-PAGE gels, served as an indicator of the maturation state of CFTR.

Immunoblot results revealed that no mature band was detected for ΔF508 CFTR, V232D CFTR,

and H1085R CFTR (Figure 6A, B, C, and D). In addition, wildtype (Figure 6A, B, C, and D),

60

Figure 5 Model of CFTR. The locations of the nucleotide-binding domains (NBD1 (light pink);

NBD2 (dark pink)) and transmembrane domains (TMD1 (light green); TMD2 (dark green)) and

the R domain (red) of CFTR are shown. Also indicated are the locations of residues V232,

H1085, and F508 in TMD1, TMD2, and NBD1 of CFTR, respectively. The structure was

generated and viewed using the PyMOL Molecular Graphics System (DeLano, 2002), which is

based on the theoretical model of CFTR structure proposed by Serohijos et al. (2008).

61

I344R (Figure 6A), S912R (Figure 6B), Y913R (Figure 6B), Y914R (Figure 6B), and Q1144R

(Figure 6C) CFTR matured as expected. However, none of the double mutants tested showed any

significant maturation, which means that none of the I344R, S912R, Y913R, Y914R, or Q1144R

mutations rescued ΔF508 CFTR, V232D CFTR, and H1085R CFTR by promoting maturation

(Figure 6A, B, and C). In other words, none of the arginine mutations acted as a suppressor

mutation for ΔF508 CFTR, V232D CFTR, and H1085R CFTR.

To generate positive controls for this experiment, V510D, I539T, and R1070W, which are

suppressor mutations known to rescue ΔF508 CFTR (Reviewed by Schmidt et al., 2011), were

introduced into ΔF508 CFTR, V232D CFTR, or H1085R CFTR cDNAs by site-directed

mutagenesis. HEK-293 cells were transfected with the cDNAs, and whole cell extracts were

subjected to immunoblot analysis using 6.5% (w/v) acrylamide gels and a CFTR polyclonal

antibody. Immunoblot results (Figure 6D) showed that when introduced into ΔF508 CFTR,

V510D, I539T, and R1070W promoted maturation of the protein. This finding is consistent with

previous studies that have demonstrated the introduction of V510D (Loo et al., 2010) or I539T

(DeCarvalho et al., 2002) in NBD1, or R1070W (Thibodeau et al., 2010) in ICL4 of ΔF508

CFTR partially corrects the folding defects to promote maturation and stability at the cell surface.

Furthermore, it was found that V510D promoted the maturation of V232D CFTR and H1085R

CFTR (Figure 6D).

62

Figure 6 Immunoblot analysis of the double mutants generated to test for suppressor

mutations. Arginine mutagenesis and immunoblot analysis were performed to test whether

I344R (A), S912R (B), Y913R (B), Y914R (B), or Q1144R (C) could rescue ΔF508, V232D, or

H1085R CFTR. V510D, I539T, and R1070W, which are known suppressor mutations were

included as positive controls (D).The arginine mutations were introduced into ΔF508 CFTR,

V232D CFTR, or H1085R CFTR cDNAs by site-directed mutagenesis, and HEK-293 cells were

transfected with the cDNAs. Whole cell extracts were subjected to immunoblot analysis using a

polyclonal antibody against CFTR. The positions of the mature CFTR (190 kDa) and immature

CFTR (170 kDa) are indicated.

63

3.2 DIRECT RESCUE USING CORRECTORS1

3.2.1 IDENTIFYING CORRECTORS THAT SPECIFICALLY RESCUE CFTR

PROCESSING MUTANTS

ΔF508 CFTR is partially functional when trafficked to the plasma membrane (Dalemans

et al., 1991). Therefore, a possible treatment for CF patients with the ΔF508 mutation would be

to promote maturation and trafficking of ΔF508 CFTR using pharmacological chaperones, which

are correctors that are specific for CFTR and are predicted to promote maturation by interacting

directly with the misfolded protein. Compared to indirect rescue approaches like low temperature

rescue, the use of pharmacological chaperones is hypothesized to yield a more stable protein, and

to cause fewer side effects by not interacting or changing the expression or activity of other

proteins. Furthermore, since most correctors identified to date require relatively high

concentrations for effective rescue in CF patients (e.g. daily doses of 100-200mg for VX-809)

(Clancy et al., 2011), ideal correctors should not be substrates of drug pumps such as P-gp,

which are predicted to reduce the bioavailability of the corrector by pumping it out of the cells.

Therefore, in this thesis, we identified correctors that are not substrates of P-gp. Previous studies

done in our lab have shown that substrates of P-gp can increase the expression of the protein by

interacting with it and promoting folding (Loo and Clarke, 1997). Furthermore, we identified

correctors that could promote maturation of ΔF508 CFTR into a more stable protein compared to

low temperature rescue.

The P-gp processing mutant G268V and the CFTR processing mutants H1085R, V232D,

and ΔF508 were used to test the specificity of various correctors - corr-5a (identified by

Pedemonte et al., 2005), corr-5c (identified by Pedemonte et al., 2005), RDR1 (identified by

1 This work constituted a publication in Biochem. Pharmacol. 83: 345-354.

64

Sampson et al. 2011) and VX-809 (identified by Van Goor et al., 2011) (Figure 7). VX-809 was

tested because it is the best corrector for CFTR identified to date (Kalid et al., 2010). Correctors

5a, 5c, and RDR1 were included because these pharmacological chaperones have been suggested

to interact directly with CFTR. Differential scanning fluorimetry studies have shown that RDR1

directly interacts with NBD1 of CFTR (Sampson et al., 2011). It is not clear which domain(s) of

CFTR corr-5a and corr-5c interact with, but the finding that the cyanoquinoline class of

compounds exhibit both corrector and potentiator activities (Phuan et al., 2011) suggest that 5a

and 5c function by direct binding to CFTR. Cyclosporin A and verapamil were included as

control compounds, as they are drug substrates of P-gp that are known to promote maturation of

P-gp but not CFTR processing mutants (Loo and Clarke, 1997). G268V P-gp was chosen

because it shows many common properties of all P-gp processing mutants, such as disrupted

domain-domain interactions and improper packing of the transmembrane segments. ΔF508,

V232D, and H1085R CFTRs were chosen because they are processing mutations located in

different domains of CFTR – V232D in TMD1, H1085R in TMD2, and ΔF508 in NBD1 (results

for ΔF508 and V232D CFTRs not shown here, see Loo et al., 2012). Furthermore, it has been

shown that the folding defects of V232D, H1085R, and ΔF508 can be repaired by the chemical

chaperones corr-4a (Caldwell et al., 2011), CFcor-325 (Loo et al., 2006), and VX-809 (Loo et al.,

2012), respectively.

65

Figure 7 Structure of correctors. This figure was reprinted from Loo, T.W., Bartlett, M.C., Shi,

L., and Clarke, D.M. (2012) Corrector-mediated rescue of misprocessed CFTR mutants can be

reduced by the P-glycoprotein drug pump. Biochem. Pharmacol. 83: 345-354.

66

To identify correctors that were specific for CFTR, we used a transient expression system

with HEK-293 cells. HEK-293 cells were transiently transfected with plasmids encoding CFTR

or P-gp processing mutants. The medium was changed 5 hours later to plain medium (control) or

medium containing a CFTR corrector at a concentration reported to rescue CFTR processing

mutants. After the cells were grown overnight to allow for expression of the protein, the cells

were harvested, and whole cell extracts were subjected to immunoblot analysis to test for

maturation (see Methods). Maturation was monitored by a shift in size due to the addition of

complex carbohydrate in the Golgi. Western blot results showed that correctors corr-5a, corr-5c,

and VX-809 promoted maturation of the H1085R CFTR processing mutant (Figure 8A). Little

mature protein band (170 kDa for G268V P-gp and 190 kDa for H1085R CFTR) was detected

when the cells were treated with plain media (i.e. < 5% of the total protein was present as mature

for untreated G268V P-gp and H1085R CFTR). It was observed that 22%, 13%, and 37% of the

total CFTR protein was present as mature when H1085R CFTR was expressed in the presence of

corr-5a, corr-5c, and VX-809, respectively (Figure 8A). Corr-5a, corr-5c, and VX-809 did not

rescue G268V P-glycoprotein (i.e. > 95% immature band and < 5% mature band with and

without corrector) (Figure 8B).

Class-5 correctors (corr-5a and corr-5c), VX-809, and RDR1 were observed to

specifically rescue CFTR processing mutants (RDR1 results not shown here, see Loo et al.,

2012), as they did not promote rescue of G268V P-gp, and thus we further tested to see whether

corr-5a, VX-809, and RDR1 promoted the stability of ΔF508 CFTR. To test the effects of the

correctors on ΔF508 CFTR stability, transfected cells were incubated overnight at 30oC in the

presence or absence of corrector to promote maturation, after which cycloheximide was added to

stop protein synthesis and the cells were moved to 37C. Cells were harvested 0, 0.5, 1, 2, 4, 8,

67

Figure 8 Effect of correctors on H1085R CFTR (A) and G268V P-gp (B). HEK-293 cells

were transiently transfected with plasmids encoding H1085R CFTR or G268V P-gp. H1085R

CFTR (A) or G268V P-gp (B) were expressed in the absence (Plain media) or presence of CFTR

correctors or P-gp substrates (verapamil, cyclosporine A) which were included as control

compounds, and whole cell extracts were subjected to immunoblot analysis. The positions of the

mature CFTR (190 kDa), immature CFTR (170 kDa), mature P-gp (170 kDa), and immature P-

gp (150 kDa) are indicated.

68

and 24 hours after addition of cycloheximide, and whole cell extracts were subjected to

immunoblot analysis using 6.5% (w/v) acrylamide gels and monoclonal antibody A52. The

mature protein bands were quantified and compared to the amount of mature protein at time 0

tocalculate the half-life of the protein. It was observed that ΔF508 CFTR had a longer half-life

when expressed in the presence of corr-5a and RDR1. Furthermore, the corrector corr-5a

stabilized both mature and immature ΔF508 CFTR (Figure 9). In the absence of correctors, the

half-lives of mature and immature ΔF508 CFTR were approximately 1 h and 0.5 h, respectively.

The half-lives of mature ΔF508 CFTR in the presence of corr-5a and RDR1 were approximately

10 h and 4 h, respectively. Compared to corr-5a and RDR1, VX-809 only had little effect on the

half-life of ΔF508 CFTR (1.3 h in the presence of VX-809). The results in Figure 9 show that

VX-809 was the most efficient corrector in promoting maturation of ΔF508 CFTR whereas

RDR1 and corr-5a were the most efficient correctors to increase stability of the mature form of

the mutant.

We also conducted cell surface labeling experiments to determine whether corr-5a

affected the trafficking of ΔF508 CFTR to the cell surface. BHK cells expressing ΔF508 CFTR

were incubated overnight in the presence or absence of corr-5a to promote maturation of the

protein. CFTR protein at the cell surface was identified by biotinylation (see Methods). ΔF508

CFTR at the cell surface was only detectable in the presence of corr-5a, suggesting corr-5a

indeed promoted trafficking of mature ΔF508 CFTR to the cell surface (Figure 10).

69

Figure 9 Stability of ΔF508 CFTR in the presence or absence of correctors. HEK-293 cells

were transiently transfected with ΔF508 CFTR. A52-tagged ΔF508 CFTR was expressed at 30C

without corrector, or in the presence of corr-5a, RDR1, or VX-809. Cycloheximide was added to

stop protein synthesis. Cells were harvested at times 0, 0.5, 1, 2, 4, 8, and 16 hours after the

addition of cycloheximide, and whole cell extracts were subjected to immunoblot analysis. The

positions of the mature ΔF508 CFTR (190 kDa) and immature ΔF508 CFTR (170 kDa) are

indicated.

70

Figure 10 Effect of corr-5a on expression of ΔF508 CFTR on the cell surface. Cell surface

labeling was performed on untransfected cells or cells expressing ΔF508 CFTR in the presence

or absence (untreated) of 20µM corr-5a. Cells were incubated overnight at 30C in the absence

of presence of corr-5a to promote maturation of the protein, after which cell surface labeling was

performed (see Methods). The position of mature ΔF508 CFTR (190kDa) is indicated.

71

3.2.2 IDENTIFYING SITES OF CORRECTOR INTERACTIONS

It was observed that corr-5a, corr-5c, VX-809, and RDR1 specifically rescued CFTR

processing mutants (RDR1 results not shown here, see Loo et al., 2012). We predicted that

correctors rescue CFTR processing mutants by binding directly to the protein, and were

interested in identifying their sites of interactions. CFTR half- molecules or domains expressed

as separate proteins were constructed for the purposes of determining the minimum CFTR

structure required for maturation and identifying sites of corrector interactions. VX-809 is the

best corrector known to date and it was found that it specifically rescues CFTR processing

mutants, and thus, it was used to identify sites of corrector interactions.

CFTR half-molecules or domains expressed as separate proteins may be useful to

determine the minimum CFTR structure required for maturation and to map the sites of corrector

interactions. For example, if a CFTR truncation mutant lacking the R domain matures in the

presence of VX-809, then it suggests that the R domain is not required for VX-809 rescue. To

map the sites of corrector interactions, the effect of VX-809 on processing of the truncation

mutants, either changes in maturation, stability, or core-glycosylation, was monitored. The

ability of VX-809 to interact with truncation mutants such as TMD1+2 lacking the NBD1

domain containing the di-acidic motif (residues 563-567) required for COPII-mediated ER

export (Wang et al., 2004) could not be examined by testing for maturation. Therefore, the

effects of VX-809 on truncation mutants lacking NBD1 were tested by assays for stability or

core-glycosylation. For instance, if VX-809 is shown to promote stability or core-glycosylation

of TMD1 and TMD2 lacking both NBDs and the R domain, then it suggests that VX-809 may

functions by interacting directly with the TMDs. To test the effect of correctors on the

maturation of CFTR truncation mutants, HEK-293 cells were transiently transfected with

72

plasmids encoding half-molecules or domains of CFTR (N-half, C-half, NBD1, TMD1, TMD2,

TMD1+2, ΔNBD1, ΔNBD2, or ΔR). The medium was changed 5 hours later to medium

containing VX-809. Cells were grown overnight at 37C to allow for expression of the half

molecule or domain, and cells were harvested on the next day. Whole cell extracts of A52-tagged

CFTR half molecule or domain were subjected to immunoblot analysis to test for changes in

glycosylation state. The glycosylation state of the two glycosylation sites located in the

extracellular loop connecting TM segments 7 and 8 of CFTR, monitored by a difference in

mobility of SDS-PAGE gels, served as an indicator of the maturation state of CFTR.

VX-809 was tested for its effects on ΔNBD2 CFTR (a CFTR truncation mutant lacking

NBD2). It was observed VX-809 rescued the CFTR truncation mutant lacking the NBD2 domain

by promoting maturation (Figure 11). ΔNBD2 CFTR was able to mature in the presence of VX-

809 despite that it lacked the NBD2 domain, suggesting the NBD2 domain was not required for

maturation of the protein, and correctors such as VX-809 do not function by binding to the

NBD2 domain.

To test whether the R domain is required for maturation and whether it contains VX-809

interaction sites, VX-809 was tested for its effects on the coexpression of C-half molecule

containing no R domain (i.e. only NBD2 and TMD2 are present) and N-half molecules

containing TMD1 and NBD1. It has been shown that complex glycosylation of the extracellular

loop between TM segments 7 and 8 is inefficient when C-half CFTR containing NBD2, TMD2,

and the R domain is synthesized alone in cells, but coexpression of C-half CFTR with N-half

CFTR containing NBD1 and TMD1 greatly improves the efficiency of complex glycosylation of

73

Figure 11 Effect of VX-809 on maturation of ΔNBD2 (Δ1197-1480) CFTR. (A) Schematic

model of CFTR. NBD2 (amino acid residues 1197-1480) is shown in red, while the rest of the

protein is shown in green. The structure was generated and viewed using the PyMOL Molecular

Graphics System (DeLano, 2002), which is based on the theoretical model of CFTR structure

proposed by Serohijos et al. (2008). (B) Samples of cells expressing ΔNBD2 CFTR and grown

in the absence (-) or presence (+) of 10µM VX-809 overnight were subjected to immunoblot

analysis. The positions of mature and immature ΔNBD2 CFTR and GAPDH (loading control)

are indicated. Untransfected cells were included as a negative control.

74

C-half CFTR. Moreover, the efficiency of complex glycosylation of C-half could be improved

even further by coexpressing C-half and N-half in the presence of VX-809 (Loo et al., 2009).

TM segments 5 and 6 of the N-half CFTR can interact with TM segments 7 and 8 of the C-half

CFTR and facilitates glycosylation by helping to orient the glycosylation sites in the extracellular

loop connecting TM segments 7 and 8 in a position that can be easily modified by oligosaccharyl

transferase. Therefore, co-expression of C-half molecules with R domain and N-half molecules

in the absence or presence of VX-809 was used as a positive control. As expected, no complex

glycosylated mature C-half band was observed for cells transfected with C-half CFTR lacking

the R domain alone, as C-half CFTR alone cannot undergo efficient glycosylation. Surprisingly,

immunoblot results showed that maturation of the C-half molecule with no R domain was

defective when coexpressed with the wildtype N-half molecule, even in the presence of VX-809

(Figure 12). We suspected this might be due to low stability without the R domain, and therefore,

we introduced V510D and Δ404-435 mutations into the N-half molecule and tested to see

whether they could enhance complex glycosylation of the C-half when coexpressed. V510D is a

known suppressor mutation for ΔF508 CFTR that promotes maturation and increases stability of

the protein when introduced into NBD1 (Loo et al., 2010). Amino acid residues 404-435

compose a region of NBD1 referred to as the regulatory insertion loop. Previous studies have

shown that removal of the regulatory insertion loop restores maturation and stability of ΔF508

CFTR (Aleksandrov et al., 2010). Both the V510D substitution and the deletion of amino acids

404-435 stabilized the protein. It was observed that coexpression of the C-half molecule lacking

the R domain and wildtype N-half molecule containing either or both of V510D and Δ404-435 in

the presence of VX-809 increased both the amount of mature protein and the amount of

75

Figure 12 Coexpression of C-half and N-half CFTRs. (A) Schematic model of CFTR. N-half

(amino acids 1-633, containing NBD1 and TMD1), C-half (amino acids 837-1480, containing

NBD2 and TMD2), and the R domain are shown in blue, green, and yellow, respectively. The

positions of the deletion of the regulatory insertion loop (amino acids 404-435) and the V510D

mutation are indicated. The structure was generated and viewed using the PyMOL Molecular

Graphics System (DeLano, 2002), which is based on the theoretical model of CFTR structure

proposed by Serohijos et al. (2008). (B) Samples of cells coexpressing C-half (no R domain) and

N-half (wildtype, V510D, Δ404-435, or Δ404-435/V510D) molecules and grown in the absence

(-) or presence (+) of 10µM VX-809 overnight were subjected to immunoblot analysis. The

positions of mature C-half, immature C-half, N-half and GAPDH (loading control) are indicated.

Untransfected cells were included as a negative control, and cells coexpressing C-half with the R

domain and N-half molecules (wildtype, Δ404-435, or Δ404-435/V510D) were included as

positive controls.

76

immature protein (Figure 12). Coexpression of C-half molecule with R domain and N-half

molecules was used as a positive control. As expected, the amount of mature C-half molecule

with R domain increased when it was coexpressed with N-half molecules in the presence of VX-

809 (Figure 12B). These results suggested that the R domain is not absolutely required for

maturation, but its presence increases the stability of CFTR. Furthermore, correctors such as VX-

809 do not interact with the R domain of CFTR.

To examine whether the NBD1 domain is required for maturation and whether it

contains VX-809 interaction sites, VX-809 was tested for its effect on the co-expression of C-

half molecule with R domain and N-half molecules containing truncated NBD1 domain. Once

again, coexpression of C-half molecules with R domain and full-length wildtype N-half

molecules in the absence or presence of VX-809 was used as a positive control. No mature C-

half band was detected for cells cotransfected with C-half molecule (+R domain) and N-half

molecules containing truncated NBD1 domain (Figure 13). Maturation of the C-half molecule

with R domain was defective when coexpressed with any of the N-half molecule containing

truncated NBD1 domain, even in the presence of VX-809, suggesting N-half molecules

containing truncated NBD1 domain failed to interact with the C-half to facilitate its

glycosylation and maturation (Figure 13). Therefore, NBD1 may contain a potential VX-809

binding site. A problem however was that deletions of NBD1 may have caused misfolding of the

domain to yield a misfolded protein that can no longer be rescued. In addition, some of the N-

half constructs lacked the COPII exit motif (residues 563-567) and were unable to mature.

Therefore, TMD constructs lacking the NBDs or R domain were used to test for VX-809 effects

on stability or core-glycosylation.

77

Figure 13 Coexpression of C-half CFTR and N-half CFTRs containing truncated NBD1. (A)

Schematic model of CFTR. N-half (amino acids 1-386, containing TMD1), and C-half with the

R domain (amino acids 634-1480, containing NBD2, TMD2, and the R domain) are shown in

blue and green, respectively. NBD1 (amino acids 387-646) is shown in red. The structure was

generated and viewed using the PyMOL Molecular Graphics System (DeLano, 2002), which is

based on the theoretical model of CFTR structure proposed by Serohijos et al. (2008). (B)

Samples of cells coexpressing C-half (with R domain) and N-half (containing truncated NBD1

domain) CFTRs and grown in the absence (-) or presence (+) of 10µM VX-809 overnight were

subjected to immunoblot analysis. The positions of mature C-half, immature C-half, full-length

N-half, truncated N-halves, and GAPDH (loading control) are indicated. Untransfected cells

were included as a negative control.

78

VX-809 was then tested for its effects on TMD1+2 CFTR (a CFTR truncation mutant

lacking the NBDs and the R domain). As TMD1+2 CFTR lacks the NBD1 domain containing

the di-acidic ER exit motif required for COPII-mediated ER export (Roy et al., 2010), it does not

get trafficked to the Golgi where it undergoes maturation, and thus, the effect of VX-809 on

core-glycosylation of the protein was monitored. It was observed that expression in the presence

of VX-809 enhanced glycosylation of TMD1+2 CFTR, and the majority of TMD1+2 were

synthesized as a glycosylated protein in the presence of VX-809 (Figure 14). Since core

glycosylation is a cotranslational event and the two glycosylation sites of CFTR are located in

the extracellular loop connecting TM segments 7 and 8, the finding that TMD1+2 CFTR core-

glycosylation in the ER was increased in the presence of VX-809 suggested that VX-809

enhanced the efficiency of insertion of TM segments 7 and 8 into the lipid bilayer. It can be

hypothesized that VX-809 functioned by interacting directly with the transmembrane domains of

CFTR to alter its folding while it is being synthesized in the ER.

Results from experiments described above revealed there are possible sites of

interactions between VX-809 and TMD1, TMD2, and NBD1 of CFTR. To determine if VX-809

affected folding of these domains expressed as separate polypeptides, the effects of VX-809 on

TMD1, TMD2, and NBD1 CFTRs were tested. HEK-293 cells were transiently transfected with

plasmids encoding TMD1, TMD2, or NBD1 CFTR cDNAs. Cells were grown overnight at 37C

to allow for expression of the proteins in the presence or absence or VX-809. The next day, cells

were harvested and whole cell extracts were subjected to immunoblot analysis to test for

expression. It was observed that the expression of TMD1 increased dramatically (approximately

3-fold) in the presence of VX-809 (Figure 15).

Previous studies have suggested that TM segments 5 and 6 of TMD1 interact with TM

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Figure 14 Effect of VX-809 on glycosylation of TMD1+2 CFTR. (A) Schematic model of

CFTR. TMD1 and TMD2 are shown in red and pink, respectively, while the rest of the protein is

shown in green. The structure was generated and viewed using the PyMOL Molecular Graphics

System (DeLano, 2002), which is based on the theoretical model of CFTR structure proposed by

Serohijos et al. (2008). (B) Samples of cells expressing TMD1+2 CFTR and grown in the

absence (-) or presence (+) of 10µM VX-809 overnight were subjected to immunoblot analysis.

The positions of unglycosylated and glycosylated TMD1+2 CFTR and GAPDH (loading control)

are indicated.

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Figure 15 Effects of VX-809 on TMD1, TMD2, and NBD1 CFTRs. (A) Schematic model of

CFTR. TMD1, TMD2, and NBD1 are shown in red, blue, and yellow, respectively, while the rest

of the protein is shown in green. The structure was generated and viewed using the PyMOL

Molecular Graphics System (DeLano, 2002), which is based on the theoretical model of CFTR

structure proposed by Serohijos et al. (2008). (B) Samples of cells expressing NBD1, TMD1, or

TMD2 and grown in the absence (plain) or presence of 10µM VX-809 overnight were subjected

to immunoblot analysis.

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segments 7 and 8 of TMD2 to promote glycosylation of CFTR by helping to properly orient the

glycosylation sites in the extracellular loop connecting TM segments 7 and 8 (Loo et al., 2009).

Furthermore, I have demonstrated in this thesis that VX-809 promoted core-glycosylation of

TMD1+2, and enhanced the expression of TMD1. Therefore, we predicted that VX-809

functions by increasing the stability of TMD1, and thereby enhances the interaction between

TMD1 and TMD2 to promote glycosylation. We assessed turnover of TMD1 CFTR after rescue

with VX-809 to determine whether VX-809 promoted the stability of TMD1 CFTR. Transfected

HEK-293 cells were incubated overnight in the presence or absence of VX-809 to promote

maturation. Cycloheximide (0.5mg/ml) was added the next day to stop protein synthesis, and

cells were harvested 0, 1, 2, 4, 6, 8, 16, and 24 hours after addition of cycloheximide. Whole cell

extracts were subjected to immunoblot analysis to monitor turnover of the protein. The TMD1

protein bands were quantitated and compared to the amount of mature protein at time 0 to

calculate the half-life of the protein. In the absence of VX-809, turnover of TMD1 CFTR was

rapid with a half-life of approximately 1.7 h. Corrector VX-809 increased the half-life of the

protein to approximately 6 h (Figure 16). These findings suggested that VX-809 may function by

interacting with TMD1 of CFTR to increase its stability, which in turn leads to enhanced

interaction between TMD1 and TMD2, resulting in increased glycosylation and maturation of

the protein. The results also suggest that VX-809 may promote maturation of CFTR processing

mutants like ΔF508 CFTR by promoting folding or stability of TMD1.

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Figure 16 Effect of VX-809 on TMD1 CFTR turnover. (A) HEK-293 cells expressing TMD1

CFTR in the presence or absence (Control) of 10µM VX-809 were first incubated at 30C to

allow for expression of the protein. Protein synthesis was stopped by addition of 0.5mg/ml

cycloheximide. The positions of TMD1 CFTR and GAPDH (loading control) in the

immunoblots are indicated. The amount of TMD1 CFTR at each time point was quantitated and

expressed relative to time 0 (B). The results are the mean of three experiments ± SD.

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3.2.3 EFFECT OF OTHER CF MUTATIONS ON STABILITY OF CFTR

ΔF508 CFTR exhibits lower stability and higher turnover rate at the cell surface

compared to wildtype CFTR as a result of disrupted NBD1-TMD2 interactions (Lukacs et al.,

1993). We predicted processing mutations in other domains, such as V232D in TMD1 and

H1085R in TMD2 of CFTR, would have less effect on the stability of mature CFTR than ΔF508.

To test whether other CF mutations lower the stability of CFTR, we examined the

stability of V232D and H1085R CFTRs after low-temperature rescue. Transfected HEK-293

cells were incubated overnight at 30C to promote maturation of the protein. 0.5mg/ml

cycloheximide was added the next day to stop protein synthesis, and cells were harvested 0, 1, 2,

4, 8, 16, and 32 hours after addition of cycloheximide at 37C. Whole cell extract was subjected

to immunoblot analysis to monitor turnover of the protein. It was found that the mature V232D

and H1085R CFTRs exhibited longer half-lives than ΔF508 CFTR (mature V232D and H1085R

CFTRs were detectable even after 32 hours, while mature ΔF508 CFTR was undetectable after 1

hour (Figure 17).

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Figure 17 Stability of other CFTR mutants. Degradation of wildtype, ΔF508, H1085R, and

V232D CFTRs was monitored overtime at 37C. After incubating the cells at 30C to promote

maturation of CFTR, 0.5 mg/ml cycloheximide was added to stop protein synthesis, and whole

cell extracts were subjected to immunoblot analysis after the indicated times at 37C. The

positions of mature and immature CFTR in the immunoblots are indicated.

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4 DISCUSSION

4.1 ARGININE SUPPRESSOR MUTATIONS

The ΔF508 mutation in CF patients causes defective folding and trafficking of CFTR

to the cell surface. We hypothesized that CFTR folding defects could be corrected by

introducing arginines in the transmembrane domains of the protein. Moreover, arginine-

scanning mutagenesis is a useful biochemical approach to learn about the structure of ABC

superfamily of transporter proteins, such as CFTR and P-gp. For instance, arginine-scanning

mutagenesis of the TM segments of the human P-gp G251V processing mutant have been

used to identify the residues lining the drug translocation pathway (Loo et al., 2009b). The

rationale was that arginine introduced into the lipid-facing positions would inhibit maturation,

arginine introduced into the aqueous face of the drug translocation pathway would not affect

maturation, and arginine introduced into the drug-binding pockets would mimic drug rescue

to promote maturation (Loo et al., 2009b). The orientations of all TM segments except TM3

and TM5 predicted by arginine-scanning mutagenesis were consistent with the mouse P-gp

crystal structure (Aller et al., 2009). It was found that the hydrophilic and hydrophobic faces

of TM3 and TM5 were in opposite directions for the predicted human P-gp structural model

and the mouse P-gp crystal structure. Jin et al. (2012) interpreted the arginine-scanning

mutagenesis data using their newly generated model of the human P-gp, based on the C.

elegans P-gp structure, and that they were in agreement. Their data also suggested that the

mouse structure was incorrect due to register errors in TM3 and TM5 of the mouse crystal

structure. Arginine mutagenesis can also be used to test potential interactions that exist

between residues. The homology models of human P-gp based on the mouse and C. elegans

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crystal structures have shown that there exist four intracellular loops (ICLs) connecting the

TMDs and the NBDs of the protein (Loo and Clarke, 2013). The C. elegans, but not the

mouse model, suggested the presence of a salt bridge between Glu256 and Arg276 of ICL2

that could play an important role in P-gp folding (Loo and Clarke, 2013). In silico docking

studies for the discovery of novel inhibitors of human P-gp relies on an accurate model of

human P-gp, and thus arginine mutagenesis was done to investigate the possibility of a

Glu256-Arg276 salt bridge in human P-gp. Loo and Clarke (2013) found that introducing

either E256R or R276E mutation into P-gp inhibited protein maturation, but rather

introducing both E256R and R276E mutations into P-gp simultaneously had no effect on

protein maturation and activity. These arginine mutagenesis results served as biochemical

evidence suggesting that the structure of ICL2 in human P-gp resembles that of C. elegans in

that there is a salt bridge between Glu256 and Arg276 of ICL2 which is essential for P-gp

folding (Loo and Clarke, 2013). Not only arginine mutagenesis, but mutagenesis experiments

in general are useful in gaining insight into protein structure. Site-directed mutagenesis have

been previously used to investigate the dimerization motifs of viral coat proteins (Deber et

al., 1993), the dimerization ability of the transmembrane domain of myelin protein zero

(Plotkowski et al., 2007), and the effect of introduction of non-native polar residues into the

TM segments of CFTR on protein folding and function (Choi et al., 2004; Wehbi et al.,

2008).

Arginine-scanning mutagenesis of TM6, TM8, and TM12 of wildtype CFTR showed

that introduced arginines can either partially inhibit maturation, completely inhibit

maturation, or cause little or no change in maturation. The results from arginine mutagenesis

experiments are useful in predicting the relative positions of the residues in the TMDs of

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CFTR. In general, introduced arginines that did not inhibit maturation were predicted to face

the aqueous pore while introduced arginines that inhibited maturation were predicted to face

the lipid or at an interface between TM segments inhibited maturation. For instance, I344R

(TM6), M348R (TM6), S912R (TM8), Y913R (TM8), Y914R (TM8), V915R (TM8), F916R

(TM8), and Q1144R (TM12) CFTRs showed maturation efficiencies similar to wildtype

CFTR (mature protein was the major product). These arginine mutations had little effect on

protein folding and maturation of the protein, suggesting they are pore-lining residues. Qian

et al. who used internal application of methanethiosulfonate (MTS) reagents to identify pore-

lining side chains in TM6 (El Hiani et al., 2010) and TM12 (Qian et al., 2011) found similar

results. They applied cysteine-reactive MTS reagents to the cytoplasmic side of open

channels, and used patch clamp recording to examine their accessibility to cysteine

substituted residues along the length of TM6 (El Hiani et al., 2010) and TM12 (Qian et al.,

2011) of a cysteine-less variant of CFTR. It was observed that intracellular MTS reagents

modified I344C and M348C of TM6 (El Hiani et al., 2010), and Q1144C of TM12 (Qian et

al., 2011) leading to a change in channel function detected by patch clamp recording,

suggesting these residues are pore-lining residues. Furthermore, the results of arginine-

scanning mutagenesis are in agreement with the models for CFTR generated and viewed

using PyMOL Molecular Graphics System (Figure 3D), which is based on the theoretical

model of CFTR structure proposed by Serohijos et al. (2008).

Iodide efflux assays revealed that M348R, V915R, and F916R CFTRs gave no efflux

at all. They were unable to conduct iodide ions possibly due to blockage of the aqueous pore

of CFTR caused by the mutations. Mutations I334R, S912R, Y913R, Y914R, and Q1144R

did not block the channel function of CFTR, and were each introduced into ΔF508, H1085R,

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and V232D CFTRs to test for potential suppressor mutations that could rescue the CFTR

processing mutants. However, it was found that none of these mutations were able to

promote maturation of the CFTR processing mutants. Loo et al. (2009b) conducted arginine

mutagenesis experiments on the transmembrane segments of P-glycoprotein, another

member of the ABC transporter family that is structurally similar to CFTR, and found that

that 38 of the arginine mutations introduced mimicked drug rescue to promote maturation of

the P-glycoprotein processing mutants. Arginine mutagenesis of TM6, TM8, and TM12 of

CFTR processing mutants yielded no suppressor mutation, while that of transmembrane

segments of P-glycoprotein yielded 38. This is most likely because P-glycoprotein, which

functions as a drug pump, contains mobile transmembrane segments lining the large

aqueous-filled drug translocation pathway that allow it to bind many drug substrates of

different shapes and sizes at its drug-binding sites by an induced-fit mechanism. By contrast,

CFTR, which functions as an ion channel that transports chloride and other ions across

epithelial cell membranes, is predicted to have a more rigid structure which would reduce the

ability of the TM segments to rotate to form hydrogen bonds.

4.1.1 CONCLUSIONS

Our experiments suggest that CFTR may not be able to be rescued by a drug-rescue

mechanism like P-gp, because no arginine suppressor mutations were identified. In contrast,

a number of second-site mutations (e.g. I539T, G550E, R553M/Q, R555K, and V510D)

which promote ΔF508 CFTR maturation and trafficking to the cell surface have been

discovered in NBD1 of CFTR (He et al., 2010; Loo et al., 2010). The large number of

suppressor mutations in NBD1 suggest that it may be a useful therapeutic target. A problem,

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however, is that the only corrector identified to date that binds to NBD1 (RDR1) only

rescues ΔF508-CFTR with low efficiency (Sampson et al., 2011). Our results suggest that

correctors that efficiently promote ΔF508-CFTR, like VX-809, interact with the TMDs by a

mechanism that is different from P-gp drug-rescue.

4.2 DIRECT RESCUE USING CORRECTORS

Direct rescue approaches for promoting maturation and trafficking of ΔF508 CFTR

using pharmacological chaperones are of higher therapeutic value than indirect rescue

approaches as they would cause less side effects by not affecting the expression or activities

of proteins involved in other metabolic pathways. We hypothesized that pharmacological

chaperones confer specificity to CFTR by binding directly to the protein to promote

maturation and enhance stability of the protein. Furthermore, ideal correctors should not be

substrates of drug pumps such as P-glycoprotein, which are predicted to reduce the

bioavailability of the corrector by pumping them out of the cells. In this thesis, we found

that the quinolines (corr-5a and corr-5c) and VX-809 showed specificity for rescue of CFTR

(Figure 8). They rescued H1085R CFTR specifically without promoting maturation of

G268V P-glycoprotein. However, results from experiments conducted in our lab assessing

the ATPase activity of P-glycoprotein in the presence or absence of VX-809 have shown that

VX-809 stimulated P-gp ATPase activity about 7-fold, suggesting VX-809 resembles P-gp

substrates such as colchicine in that it stimulates ATPase activity but does not have the

ability to rescue P-gp processing mutants (Loo et al., 2011). This possibly explains why

relatively high doses of VX-809 are required for effective rescue in patients with ΔF508

CFTR.

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We investigated the effects of corr-5a, RDR1, and VX-809 on the stability of ΔF508-

CFTR. VX-809, one of the most effective correctors discovered to date, had little effect on

the stability of mature ΔF508-CFTR (Figure 9). The half-lives of mature ΔF508 CFTR in the

absence and presence of VX-809 were approximately 1 h and 1.3 h, respectively. He et al.

(2012) found similar results conducting metabolic pulse-chase experiments in ΔF508-CFTR-

expressing BHK cells to examine the effect of VX-809 on the turnover of mature ΔF508-

CFTR. They demonstrated that mature ΔF508-CFTR remained short-lived with a half-life of

4.5 h after treatment with VX-809. Mature wildtype-CFTR was observed to have a half-life

of 14 h. They concluded that VX-809 is capable of improving ΔF508-CFTR maturation

without increasing the stability of the mature protein product at the cell surface. However,

Goor et al. (2011) conducted metabolic pulse-chase experiments in ΔF508-CFTR-expressing

human bronchial epithelial (HBE) cells and observed that the half-life of VX-809-corrected

ΔF508-CFTR was similar to that of wildtype CFTR (approximately 16 – 24 h). The

discrepancy among the studies might be due to the different cell lines used. He et al. (2012)

further examined the effects of VX-809 on the assembly of ΔF508-CFTR by conducting

trypsin digestion and disulfide cross-linking experiments. It was observed that the fragment

patterns of ΔF508-CFTR treated with VX-809 after trypsin digestion were similar to those of

the wildtype-CFTR, and disulfide cross-linking between domains known to occur in wildtype,

but not ΔF508 CFTR, was restored after treatment of ΔF508 CFTR with VX-809, suggesting

correctors such as VX-809 functions by promoting the proper assembly of ΔF508 CFTR. He

et al. (2012) concluded that in order to obtain more effective rescue, it is necessary to

combine correctors such as VX-809, which supports assembly, with another mechanism

which provides stabilization to the properly assembled mature ΔF508 CFTR. In support of

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this conclusion, they tested the effect of VX-809 on ΔF508-CFTR containing known NBD1-

stabilizing second-site mutations, such as V510D. It was found that each of VX-809 and

V510D alone improved ΔF508-CFTR maturation, and an additive effect on protein

maturation was observed for VX-809-corrected ΔF508/V510D-CFTR (He et al., 2012).

Other studies have also suggested completely effective correction of ΔF508-CFTR may

require a combination of correctors (Mendoza et al., 2012; Rabeh et al., 2012). Mendoza et

al. (2012) conducted computational analysis to identify residues statistically coupled to

Phe508 and examined how mutations at these positions affect NBD1 and CFTR folding.

They observed that simultaneous correction of the NBD1 folding defect and the NBD1-ICL4

interface defect by introducing a suppressor mutation that improves ΔF508 NBD1 folding

yield (I539T, R555K, G550E, or R553M), and a suppressor mutation that corrects the

NBD1-ICL4 defect into ΔF508-CFTR resulted in restoration of CFTR maturation and

function to near wildtype levels. They concluded that the deletion of Phe508 results in

misfolding of NBD1 and disruption of the NBD1-ICL4 interface, and correction of either

defect alone is insufficient to restore wildtype maturation and function. Rabeh et al. (2012)

examined the effects of introducing second-site suppressor mutations in combinations on

ΔF508 CFTR folding efficiency and cell surface expression, and reached the same

conclusion that both the ΔF508 NBD1 folding defect and NBD1-ICL4 interface defect must

be corrected to restore wildtype folding, processing, and function.

The effects of VX-809 on the maturation of CFTR-half molecules or domains

expressed as separate proteins were examined. It was found that NBD2 and R domains are

not required for the maturation of CFTR, and furthermore, TMD1, TMD2, and NBD1

possibly contain potential corrector binding sites. It was observed that VX-809 increased the

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stability of TMD1 (Figure 16A). We hypothesize that the corrector VX-809 functions by

enhancing the interaction between TMD1 and TMD2 to result in increased glycosylation and

maturation of the protein. VX-809 promoted the core-glycosylation of TMD1+2 CFTR

(Figure 14), and it enhanced the stability of mature TMD1 by about 3-fold (Figure 16B).

Okiyoneda et al. (2013) also examined the effect of VX-809 on different CFTR truncation

mutants (TMD1, TMD1+NBD1, TMD1+NBD1+R, NBD1+R+TMD2+NBD2,

R+TMD2+NBD2, NBD1+R+TMD2). Their immunoblot results showed that VX-809

increased the steady-state expression of all truncation mutants containing TMD1, and had no

effect on mutants that lacked TMD1, suggesting that VX-809 may act on TMD1 of CFTR.

Furthermore, similar to what we found with VX-809, experiments done by others in our lab

have shown that other correctors such as corr-5a, corr-5c, corr-2b, and 15Jf are also able to

enhance glycosylation of TMD1+2 CFTR (Loo et al., 2011). It is not known whether these

correctors function by directly binding to and stabilizing TMD1.

In order to understand how correctors function, it is important to take a look at the

complex CFTR biosynthetic process. Synthesis of CFTR is initiated on free ribosomes in the

cytosol, and ER targeting begins when signal recognition particle recognizes and binds an

emerging hydrophobic signal sequence near the amino terminal end of the growing

polypeptide (Reviewed by Kim and Skach, 2012). The signal recognition particle binds to its

receptor on the ER membrane, and brings the ribosome nascent chain complex to a protein-

conducting channel, the Sec61 translocon, which facilitates the translocation of hydrophilic

polypeptide segments across the endoplasmic reticulum membrane and the lateral integration

of transmembrane segments into the lipid bilayer. Alternating signal anchor and stop transfer

sequences that sequentially open and close the translocon pore determines the topology of

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the transmembrane segments (Reviewed by Kim and Skach, 2012). The topology of TM1

and TM2 of CFTR is achieved by two alternate pathways – either by a co-translational

mechanism in which translocation is initiated by TM1 and terminated by TM2 (about 25% of

nascent CFTR polypeptides), or a post-translational mechanism in which TM2 initiates

translocation and directs TM1 into the translocon (about 75% of nascent CFTR polypeptides)

(Reviewed by Kim and Skach, 2012). TM3 has been suggested to emerge from the ribosome

exit tunnel into the translocon after topology of TM1 and TM2 is established, as the loop

between TM2 and TM3, the first intracellular loop (ICL1), is about 56 residues in length

(Reviewed by Kim and Skach, 2012). TM3 and TM4 are very closely spaced – they are

connected by the second extracellular loop (ECL2), which is only 5 residues, and moreover,

the weak signal anchor activities of each of TM3 and TM4 can only efficiently initiate ECL2

translocation when both TMs are present simultaneously (Sadlish and Skach, 2004). Thus, it

has been suggested that TM3 and TM4 enters the translocon at the same time with a helical

hairpin configuration. A similar paired translocation mechanism has been proposed for TM5

and TM6, TM9 and TM10, and TM11 and TM12, as the loops between them are short in

length – 1, 2, and 5 residues, respectively (Sadlish and Skach, 2004). CFTR contains two N-

linked glycosylation sites in the extracellular loop between TM segments 7 and 8. When the

glycosylation consensus sequence Asn-X-Ser/Thr, where X is any amino acid except proline,

is at least 12 to 14 residues from the ER membrane, the oligosaccharyltransferase complex

(OST complex), which contains a 50kDa protein capable of catalyzing the transfer of the

oligosaccharide, binds to the nascent polypeptide and catalyzes the transfer of a

(Glucose)3(Mannose)9(N-acetylglucosamine)2 group from a dolichol pyrophosphate donor to

the Asn residue (Nilsson and von Heijne, 1993). N-linked glycosylation is crucial for proper

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folding and cell surface expression of CFTR (Glozman et al., 2009). Correctors, such as VX-

809, may function by enhancing the efficiency of insertion of TM segments 7 and 8 of C-half

CFTR into the lipid bilayer, and thereby allow for proper glycosylation of the glycosylation

sites between TM7 and TM8 to occur. This is supported by results from our experiments

indicating that expression in the presence of VX-809 enhanced glycosylation of TMD1+2

CFTR, and the majority of TMD1+2 were synthesized as a glycosylated protein in the

presence of VX-809 (Figure 14). It must be noted however, that there may be more subtle

effects on full-length CFTR processing mutants since they are efficiently core-glycosylated.

Our experiments suggested that TMD1, TMD2, and NBD1 are potential targets to

develop new correctors. Okiyoneda et al. (2013) performed planar phospholipid bilayer

studies to show direct interactions of VX-809 with CFTR. They showed that the opening

probability of temperature-rescued ΔF508 CFTR reconstituted in an artificial planar

phospholipid bilayer decreased dramatically when the temperature was raised from 24C to

36C, but in the presence of VX-809, the opening probability of the mutant channel

remained the same after raising the temperature. Their experiment provided evidence

suggesting that VX-809 can directly interact with CFTR. Moreover, previous studies

conducted by others have provided evidence suggesting that correctors such as RDR1, VX-

532, corr-4a, corr-2b, and corr-5a and corr-5c, act as pharmacological chaperones for ΔF508

CFTR by directly interacting with TMD1, TMD2, or NBD1 of the protein. Sampson et al.

(2011) conducted differential scanning fluorimetry to show that RDR1 directly interacts with

NBD1 of CFTR. Furthermore, it was found in our lab that when NBD1 ΔF508 CFTR was

expressed in the presence of RDR1 its stability increased by about 3-fold (Loo et al., 2011).

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The corrector VX-532 has been predicted to interact with the NBD1-TMD2 interface of

CFTR (Wellhauser et al., 2009). Results from experiments examining the effect of VX-532

on the stability of NBD1 have shown that it was able to stabilize NBD1 at high

concentrations (Loo et al., 2011). Grove et al. (2011) investigated the mechanism of corr-4a,

another bisaminomethylbithiazole corrector like 15Jf, and found that it interacts with and

stabilizes TMD2. Corr-2b related arylaminothiazoles (Pedemonte et al., 2011) and

cyanoquinoline class of compounds (Phuan et al., 2011) exhibit both corrector and

potentiator activities. This serves as evidence suggesting corr-2b and corr-5a function by

direct binding to CFTR, although which domain(s) of the protein they interact with has not

been investigated.

The ΔF508 mutation, located in NBD1, causes misfolding of NBD1 (Thibodeau et al.,

2004) and disrupts NBD1-TMD2 interactions (Lewis et al., 2004). We were interested in

testing whether processing mutations in other domains, such as V232D in TMD1 and

H1085R in TMD2 of CFTR, would have less effect on the stability of mature CFTR than

ΔF508. Our experiments suggested our prediction to be correct. Mature V232D and H1085R

CFTRs exhibited half-lives that were at least 10-fold longer than ΔF508 CFTR (Figure 17).

A possible explanation for this observation is that the ΔF508 mutation results in misfolding

of the NBD1 domain (Thibodeau et al., 2004) and causes defective domain-domain

interactions (Chen et al., 2004), while V232D and H1085R mutations have more localized

effects on protein folding in the TMD1 and TMD2 domains, respectively. Therien et al.

(2001) have demonstrated that the V232D mutation in TM4 leads to formation of a hydrogen

bond between Asp-232 and Gln-207, which changes the orientation of TM3 relative to TM4

and results in improper packing of TMD1. The H1085R mutation may cause localized

96

folding defects in TMD2.

4.2.1 CONCLUSIONS

In summary, we identified in this thesis a number of correctors which were able to

rescue CFTR processing mutants without promoting maturation of P-glycoprotein mutants.

These correctors have been suggested, either in this thesis or in previous studies done by

others, to rescue ΔF508 CFTR, possibly by directly interacting with TMD1, TMD2, or

NBD1 of the protein. More direct binding assays like the one done by Okiyoneda et al. (2013)

should be performed in the future to confirm direct corrector interactions with CFTR.

Furthermore, CF drug therapy should aim to develop compounds resembling these correctors

which show specificity for CFTR and function by acting on TMD1, TMD2, or NBD1 of

CFTR. TMD1 may be an attractive target to promote maturation since it appears to contain a

VX-809 binding site. Stabilization of ΔF508-CFTR may require the use of an additional

corrector that binds to NBD1.

4.3 FUTURE DIRECTIONS

It was found in this thesis that TMD1 appears to contain a VX-809 binding site, and

thus, it may be an attractive target to promote maturation of CFTR. In order to confirm the

suspected interaction between TMD1 of CFTR and the corrector VX-809, it is necessary to

assay for any direct interactions between TMD1 and VX-809 using NMR spectroscopy or in

vitro binding assays. Furthermore, it was found in this thesis that VX-809 had little effect on

the stability of mature ΔF508-CFTR (Figure 9). Other studies have suggested completely

effective correction of ΔF508-CFTR may require a combination of correctors – the deletion

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of Phe508 results in misfolding of NBD1 and disruption of the NBD1-ICL4 interface, and

correction of either defect alone is insufficient to restore wildtype maturation and function

(Mendoza et al., 2012; Rabeh et al., 2012). Therefore, while VX-809 promotes the proper

assembly of ΔF508 CFTR (He et al. 2012), stabilization of ΔF508 CFTR may require the use

of an additional corrector that binds to and stabilizes NBD1. To make cystic fibrosis

pharmacological therapy more feasible for CF patients, we should aim to identify and

develop correctors that targets NBD1, possibly utilizing high-throughput screening assays or

ΔF508-CFTR cell-based functional or biochemical assays.

98

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Stone, A.J., Olson, E.R., Ordonez, C.L., Campbell, P.W., Ashlock, M.A., and Ramsey, B.W.

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