2. ii
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.
3. iii
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.
4. iv
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.
5. v
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
6. vi
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
7. vii
LIST OF TABLES
Table 1 Classes of CFTR Mutations that cause cystic fibrosis.........................................19
8. viii
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
9. ix
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
10. 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
11. 2
(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)
12. 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
13. 4
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
14. 5
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
15. 6
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
16. 7
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
17. 8
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
18. 9
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).
19. 10
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
20. 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,
21. 12
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
22. 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-
32
P]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
23. 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
24. 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
25. 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
26. 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.
27. 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).
28. 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
29. 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
30. 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
31. 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
32. 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,
33. 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
34. 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.
35. 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
36. 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
37. 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
38. 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
39. 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
40. 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
41. 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
42. 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
43. 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).
44. 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.
45. 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
46. 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.
47. 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
48. 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.
49. 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),
50. 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
51. 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
52. 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
53. 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
54. 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.
55. 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
56. 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.
57. 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.
58. 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
59. 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
60. 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).
