FXYD5 MODULATES NA,K-ATPASE ACTIVITY AND IS
INCREASED IN CYSTIC FIBROSIS AIRWAY EPITHELIA
By
TIMOTHY J. MILLER
Submitted in partial fulfillment of the requirements for the
Degree of Doctor of Philosophy
Dissertation Advisor: Dr. Pamela B. Davis
Department of Pharmacology
CASE WESTERN RESERVE UNIVERSITY
May, 2008
Case Western Reserve University
School of Graduate Studies
We hereby approve the dissertation of
Timothy J. Miller
candidate for the Ph.D degree *.
(signed) Michael Maguire (chair of the committee)
Pamela Davis
Mitch Drumm
Ruth E. Siegel
______
Date March 21, 2008
* We also certify that written approval has been obtained for any proprietary material contained therein.
ii
DEDICATION
The achievement of any worthwhile goal is often accomplished through hard work and commitment. It is often easier to succor such accolades with the inspiration of a beautiful wife and son.
For her love, patience and guidance throughout our journey, her supremely generous and kind nature, and the joy of my life,
I dedicate this thesis to my wife, Molly Megan Gallogly, and my son, Gavin Bryce
Miller.
iii
TABLE OF CONTENTS
LIST OF TABLES……………………………………………………………………. vii
LIST OF FIGURES………………………………………………………………….. viii
LIST OF ABBREVIATIONS…………………………………………………………. ix
ACKNOWLEDGEMENTS……………………………………………………...……. xi
ABSTRACT………………………………………………………………………….. xiv
CHAPTER 1: BACKGROUND…………………………………………………….….1 Cystic Fibrosis is caused by defective ion transport……………………..….1 Na+ hyperabsorption and airway surface liquid dehydration…………….....3 The contribution of the Na,K-ATPase………………………………………...6 The FXYD family: modulators of Na,K-ATPase activity………………….....8 FXYD5…………………………………………………………………………..11
CHAPTER 2: CHARACTERIZATION OF HUMAN AND MOUSE FXYD5….….18 Abstract………………………………………………………………………....18 Introduction……………………………………………………………………..19 Methods…………………………………………………………………………21 Results………………………………………………………………………….30 A comparison of human and mouse expression profiles………….30 Analysis of human and mouse FXYD5……………………………...33 Discussion………………………………………………………………………36
CHAPTER 3: FXYD5 MODULATES NA,K-ATPASE ACTIVITY AND IS INCREASED IN CF AIRWAY EPITHELIA…………………………………………58
Abstract……………………………………………………………………..…..58 Introduction………………………………………………………………….….60 Methods…………………………………………………………………………62 Results………………………………………………………………………….73 Flag-FXYD5 is expressed at the cell membrane…………………...73
iv FXYD5 alters Na,K-ATPase pump kinetics…………………………74 FXYD5 is downregulated after ENaC activation……………………75 FXYD5 is decreased in the lungs of Scnn1b transgenic mice……76 FXYD5 is upregulated in the nasal epithelia of CF mice………….76 FXYD5 is increased in lungs of CF mice……………………………77 CFTR inhibition upregulates FXYD5 in human epithelia…………..78 Discussion………………………………………………………………………80
CHAPTER 3 ADDENDUM: FXYD5 IS INCREASED IN THE LUNGS OF CF MICE AFTER INFECTION WITH P. AERUGINOSA………………..102
Introduction……………………………………………………………………102 Methods……………………………………………………………………….105 Results and Discussion……………………………………………………...109
CHAPTER 4: S163 IS CRITICAL FOR FXYD5 MODULATED WOUND HEALING IN AIRWAY EPITHELIA………………………………………………..113
Abstract………………………………………………………………………..113 Introduction……………………………………………………………………114 Methods……………………………………………………………………….116 Results………………………………………………………………………...121 Mutations in Ser163 alter FXYD5 cellular localization……………121 S163 mutations alter FXYD5/Na,K-ATPase interaction………….122 S163 modulates wound healing in murine airway epithelia……..123 Discussion…………………………………………………………………….124
CHAPTER 5: SUMMARY, CONCLUSIONS AND FUTURE DIRECTIONS…..135
Summary………………………………………………………………………135 FXYD5 modulates Na,K-ATPase pump affinity for Na+ and K+....136 Consequences of increased FXYD5 expression in the airway….139 Secondary effects of FXYD5/Na,K-ATPase interaction………….143 Pro-inflammatory signals upregulate FXYD5 expression………..148 Future directions……………………………………………………………...151
v
APPENDICES………………………………………………………………………..161
Neighbor joining alignment of known FXYD5 sequences………..……...161 Human and mouse primer sequences……………………………………..162 Future directions materials and methods………………………………….163
REFERENCE LIST…………………………………………………………………..169
vi LIST OF TABLES
Table 2-1. Comparison of human and mouse FXYD5………………………….…41
Table 3-1. Comparison of intracellular Na+ and extracellular K+ activation of Na,K-ATPase pump activity in FXYD5 transfected MDCK cells...... 89
LIST OF FIGURES
Figure 1-1. Post-Albers Na,K-ATPase transport scheme…………………………16 Figure 1-2. Human FXYD5 cDNA and protein sequence…………………………17 Figure 2-1. Unigene profile of human FXYD5 tissue expression………………...43 Figure 2-2. Unigene profile of murine Fxyd5 tissue expression………………….46 Figure 2-3. Tissue distribution of murine Fxyd5 expression….…………………..48 Figure 2-4. Fxyd5 is highly expressed in murine lung tissue……………………..49 Figure 2-5. FXYD5 is expressed is multiple cell lines……………………………..50 Figure 2-6. Antibody 562 is specific for murine FXYD5 …………..………………51 Figure 2-7. FXYD5 is strongly expressed in murine epidermis …..……………...53 Figure 2-8. Murine FXYD5 tissue expression ………..…………………………….54 Figure 2-9. Alignment of human and mouse FXYD5………………………………55 Figure 2-10. Comparison of human and murine FXYD5 chimeras……….……...57 Figure 3-1. Flag-FXYD5 is membrane localized in MDCK cells...... 86 Figure 3-2. FXYD5 modulates Na,K-ATPase activity...... 88 Figure 3-3. ENaC activation increases Na,K-ATPase but decreases FXYD5.....91 Figure 3-4. FXYD5 is decreased in lungs from Scnn1b overexpressing mice.....93 Figure 3-5. FXYD5 is upregulated in the nasal epithelia of S489X-/- CF mice…95 Figure 3-6. FXYD5 is increased in airway epithelia of CF mice…………………..97 Figure 3-7. CFTR inhibition upregulates FXYD5 in human airway epithelia…….99 Figure 3-8. Model of FXYD5 role in Na+ absorption in CF airway epithelia……101 Figure 3-9. Fxyd5 is increased in CF mouse lungs after bacterial infection…...112 Figure 4-1. Immunoblot of FXYD5-Flag in HEK 293 and LA4 cells…………….128 Figure 4-2. S163 mutations alter FXYD5 membrane localization………………130 Figure 4-3. Ser163 modulates FXYD5 interaction with Na,K-ATPase……..…..132 Figure 4-4. FXYD5 modulates wound repair in murine airway epithelial cells...134 Figure 5-1. Regulation and role of FXYD5 in ASL dehydration...... 142 Figure 5-2. Helical wheel plot of human FXYD5 transmembrane domain……..146 Figure 5-3. FXYD5 S170 is an SGK1 consensus phosphorylation site………..158 Figure 5-4. SGK1 phosphorylates FXYD5 S170………………………………….159 Figure 5-5. FXYD5 S170 mutations affect cell localization in MDCK cells…….160
viii LIST OF ABBREVIATIONS a.a.: amino acid
ASL: airway surface liquid
ATP: adenosine triphosphate
BALF: broncheoalveolar lavage fluid
Bp: base pair
Ca+2: calcium
CCL2: chemokine (C-C motif) ligand 2
CCR2: chemokine (C-C motif) receptor 2
CF: cystic fibrosis
CFTR: cystic fibrosis transmembrane conductance regulator
CHIF: corticosteroid hormone-induced factor
COPD: chronic obstructive pulmonary disorder
DDD: digital differential display
ENaC: epithelial sodium channel
EST: expressed sequence tag
FXYD: phenylalanine-X-tyrosine-aspartic acid
HEK293: human embryonic kidney cells 293
Kb: kilobase kDa: kilodalton
MDCK: madin-darby kidney cells
MCC: mucociliary clearance
Na,K-ATPase: sodium, potassium adenosine triphosphatase
ix Nasal PD: nasal potential difference
NF-κB: nuclear factor kappa-B
PA: Pseudomonas aeruginosa
PCL: pericilliary layer
RAA: renin-angiontensin-aldosterone axis
RIC: Related to Ion Channel
RT: room temperature
SDS: sodium dodecyl sulfate
SGK1: serum/glucocorticoid induced kinase siRNA: silenced RNA qRT-PCR: quantitative reverse-transcription polymerase chain reaction
x Acknowledgments
The decision to go back to graduate school wasn’t easy, and wouldn’t have been possible without help from my father, Peter V. Miller. The “Miller
Foundation” is gratefully acknowledged for emotional, financial and humoral support over the past 7 years. Similarly, my mother, Wendy E. Murray, was a source of strength and determination for me and I’m proud to have had her support.
I left a wonderful workplace to join the ranks of graduate students, but it was due to the research I performed at Copernicus Therapeutics, Inc. that convinced me I wanted to work on cystic fibrosis for my thesis project. It was quite a boon to have a general field of work already chosen, and it helped propel my initial studies. I’ll always value the time I spent at Copernicus, and I thank them and Dr. Mark Cooper, M.D. for supporting my transition from employee to student.
While I may have had a general interest in CF research, this project originated from an observation made by Dr. Aura Perez, M.D., Ph.D. At the time, the FXYD family had just been identified, and FXYD5 came up on a short list of genes altered on a CF microarray. She generously allowed me to make this my project. The rest is history.
No student can perform their work alone. I was able to successfully publish my work only after collaboration with the members of the Davis lab. In particular, Yongyi Qian was a source of information for experimental design as well as a contributor to the FXYD5 project. She is a valuable asset to any lab,
xi and will be missed. Similarly, Xuguang Chen was helpful in troubleshooting a variety of experiments and was always willing to listen about my next “big experiment”. He helped provide a fun and interactive environment as well as graduate student solidarity.
Much of my thesis work was made possible as a result of the hard working staff in the Cystic Fibrosis Animal Core. My heartfelt thanks go to Alma Wilson,
Christian van Heeckeren and Veronica Peck. They were always willing to lend a hand – or a tail, as the case may be. I’d also like to acknowledge the staff of the
Pediatric Pulmonary department at Case Western and University Hospitals. In particular, Dr. Jim Chmiel, M.D., was instrumental in teaching me what it felt like to have your nose scraped, and his staff helped me acquire clinical samples for my work. I also want to acknowledge Dr. Tom Kelley, Ph.D for preparing mouse nasal epithelia for some of my studies, and thank him for his helpful discussions.
Without the help and patience of my thesis committee I wouldn’t have been able to go the distance. Dr. Mike Maguire, Ph.D gave me many opportunities to increase my skills as a teacher and lecturer and thoughtfully sent me research articles related to my work. I’ll always appreciate the advice he’s provided over the past few years, and it’s because of him that I’ll always consider myself lucky enough to “be an explorer, discovering new science”. I’d like to thank Dr. Ruth Siegel, Ph.D, for her continued enthusiasm in my project.
Whenever I talked to her about my work, she encouraged me to look at my results from a different angle. I’ve known Dr. Mitch Drumm, Ph.D, for many years, and thoughout all of them he’s been friendly, helpful, and understanding.
xii When I first became interested in academic research, Mitch was the first person who helped generate a sense of wonder in the search for a cure for CF. His gentle yet insightful approach to science has guided many graduate students before me, and I’m grateful to have had his help.
When I first joined the lab of Dr. Pamela B. Davis, M.D, Ph.D, she was the chief of the Pediatric Pulmonary division at Case Western and a founding member of Copernicus Therapeutics, Inc. We didn’t really know each other, and she took me on as a graduate student in 2001 with a particular project in mind.
When the preliminary studies of that project didn’t yield much progress, she generously allowed me to develop the “side project” I had been working on,
FXYD5, into my thesis. Since then, even as she’s risen in the ranks, Pam has treated me more of a colleague than a graduate student, an honor I’ll never forget. She’s helped make me a better scientist and writer, and I know I’ll look back fondly on the days in her lab as the Golden Days of Graduate School.
xiii FXYD5 modulates Na,K-ATPase activity and is increased in cystic fibrosis airway epithelia
Abstract
by
Timothy J. Miller
Cystic fibrosis (CF) is a common genetic disease among Caucasians, caused by lesions in the cystic fibrosis transmembrane conductance regulator gene, CFTR. This defect leads to the inability to transport chloride through the apical membrane, particularly in epithelia lining the airway. A hallmark of CF is the hyperabsorption of sodium across the airway epithelia, leading to decreased airway surface liquid volume and the inability to clear sticky, mucous plugs from the airway surface. This contributes to the smoldering inflammation and excessive growth of pathogenic bacteria typically seen in the lungs of CF patients and is often the primary cause of mortality.
Although the exact link between CFTR defects and the inability to clear trapped, pathogenic bacteria remains unclear, it has been postulated that increased sodium absorption and decreased airway surface liquid are critical modulators of disease pathogenesis in the lung. Transepithelial sodium absorption begins with entry through the epithelial sodium channel (ENaC) in the apical membrane and exits through the Na,K-ATPase in the basolateral membrane. We identified FXYD5, also known as Dysadherin, as a gene upregulated in CF airway epithelia. FXYD5 belongs to a family of tissue-specific
xiv regulators of the Na,K-ATPase, and thus we hypothesized that FXYD5 may modulate Na,K-ATPase activity and contribute to disease pathogenesis in CF.
Therefore, we determined the effects of FXYD5 overexpresison on Na,K-
ATPase activity in an epithelial cell model. FXYD5 significantly increased the apparent affinity for Na+ 2-fold, and decreased the apparent affinity for K+ by 60%
+ with a 2-fold increase in Vmax(K+), a pattern that would increase activity and Na removal from the cell. To test the effect of increased sodium uptake on FXYD5 expression, we analyzed MDCK cells stably transfected with an inducible vector expressing all three ENaC subunits. Na,K-ATPase activity increased 6-fold after
48-hour ENaC induction, but FXYD5 expression decreased 75%. FXYD5 expression was also decreased in lung epithelia from mice that overexpress
ENaC, suggesting that chronic Na+ absorption by itself downregulates epithelial
FXYD5 expression.
However, we counterintuitively found that FXYD5 was significantly increased in the lungs and nasal epithelium of CF mice as assessed by RT-PCR, immunohistochemistry and immunoblot analysis (P<0.001). FXYD5 was also upregulated in nasal scrapings from human CF patients compared to controls
(P<0.02). Treatment of human tracheal epithelial (HTE) cells with a CFTR inhibitor (CTFRinh-172) confirmed that loss of CFTR function correlated with increased FXYD5 expression (P<0.001), which was abrogated inhibitors of NF-
κB. Similarly, stimulation of NF-κB activity with the pro-inflammatory cytokines
TNFα/IL-1β upregulated FXYD5 expression and was blocked by a separate chemical inhibitor of NF-κB. We also found that overexpression of FXYD5
xv increased wound healing in airway epithelial cells, which was modulated by negative charge at S163, suggesting a role for FXYD5 in epithelial motility and regeneration in the airway. Collectively, these data show that FXYD5 is upregulated in CF epithelia and this change may exacerbate the Na+ hyperabsorption and surface liquid dehydration observed in CF airway epithelia.
xvi CHAPTER 1: BACKGROUND
Cystic fibrosis is caused by defective ion transport
Roughly 52,000 years ago, during the Paleolithic post-glacial warming epoch, mutations in a particular gene occurred in the human population living in
Asia [1]. Immigrants carried these mutations into the European population, and it is hypothesized that these mutations conferred a selective advantage in heterozygote carriers, such as the ability to resist dehydration by Vibrio cholerae or enterotoxins produced by Salmonella typhi [2, 3]. Later, some of the children born from heterozygotic parents would be recognized as having a “salty taste”, and that these children would likely soon die.
The earliest evidence referring to this salty taste came from a document written in 1606, “Das Kind sterbt bald wieder, dessen Stirne beim Kussen salzig schmeckt”∗, perhaps as a reference to cystic fibrosis [4, 5]. It wasn’t until 1938 that cystic fibrosis was identified as a disease by D.H. Andersen, who correlated the characteristic fibrotic pancreas with the intestinal obstruction and respiratory complications with what we now commonly associate with CF [6]. Respiratory tract obstruction by thick mucus, and subsequent bacterial colonization, were added to the generalized description of the disease in 1948 [7]. Soon it would be recognized that the sweat of CF patients was abnormal, containing fivefold higher concentrations of sodium and chloride [8-10], which led to the development of the pilocarpine iontophoresis test (“the sweat test”) that is still used today to diagnose CF [11, 12].
∗ The child will soon die, whose forehead tastes salty when kissed.
Despite this hint, it was unclear that this was the primary defect responsible for CF until approximately 1980 [13, 14]. Knowles and colleagues provided more direct evidence of an ion transport defect when they demonstrated electrophysiological abnormalities in the nasal potential difference of CF neonates prior to bacterial infection, noting that amiloride corrected the nasal PD
[15]. They concluded that increased sodium absorption was a hallmark of CF in airway epithelia [15, 16]. Paul Quinton demonstrated that improper chloride transport was the underlying cause of CF in sweat glands in 1983, and the gene responsible for CF was identified by three separate, colloborating groups in 1989
[17-21].
It is now known that mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) result in cystic fibrosis (CF), an autosomal- recessive genetic disease common among Caucasians [18, 19]. CFTR is a cAMP regulated chloride channel present in the apical membrane of epithelial cells. Mutations in CFTR result in inactive, improperly folded or mistargeted protein, leading to the inability to properly transport chloride. Patients with CF experience elevated sweat chloride concentration, intestinal obstructions, congenital bilateral absence of the vas deferens and pancreatic insufficiency [11].
Importantly, these patients also suffer from impaired mucociliary clearance
(MCC) in the airways, which provides a fertile environment for inhaled bacteria to adhere and flourish. Eventually, this leads to inflammation, bronchiecstasis and death from lung infection.
2 Na+ hyperabsorption and airway surface liquid dehydration
Mucociliary clearance is the innate defense mechanism for clearing particulate matter from the conducting airways and requires that the airway surface is sufficiently lubricated for efficient mechanical expulsion. The airway surface is covered in a thin layer of low-viscosity fluid that is maintained by a balance of chloride secretion and sodium absorption across the airway epithelium [22]. Proper volume maintenance is a critical factor regulating the composition and height of airway surface liquid (ASL) and is coordinately controlled through active ion transport [23, 24]. The ASL is composed of two layers - a mucus layer, consisting of heavily-glycosylated mucins that form a gel- like layer on top of the ASL, and a mucus-free zone at the cell surface, the pericilliary liquid layer (PCL) [25]. ASL volume homeostasis is disrupted in CF airway epithelia due to the failure to secrete chloride, which results in the failure to maintain normal PCL height (∼ 7 µm) and increases ASL viscosity. Combined with increased mucus production, this interferes with the normal beating of cilia
[26-28]. While decreased ASL height has been observed in CF airway epithelia, the composition and cause have been debated.
This has led to the proposal of two hypotheses to explain the reduced depth of the airway surface liquid lining the epithelia of the CF airway. The compositional hypothesis emphasizes the contribution of CFTR as the main transporter for chloride secretion, responsible for maintaining an isotonic or slightly hypotonic luminal solution in normal ASL. The salt-sensitive small molecule defensins secreted into the airway lumen are active in the low-salt ASL
3 and protect the airway much like a chemical shield [25]. This hypothesis predicts that in CF airways the ASL is hypertonic, which inappropriately inactivates antimicrobial peptides and promotes bacterial growth. However, the data indicate that both CF and non-CF ASL are nearly isotonic and suggest that airway dehydration drives mucus accumulation [23, 29, 30].
While it is currently accepted that mutant CFTR is unable to inhibit the epithelial sodium channel (ENaC), the relative contribution of sodium hyperabsorption to disease pathogenesis and severity remains controversial [31].
Early studies demonstrated that the apical membrane of CF airway epithelia was impermeable to chloride ions, but an increase in amiloride-sensitive sodium absorption through ENaC was also observed [32, 33]. ENaC is a heteromultimeric protein composed of three subunits, α, β, and γ encoded by the genes Scnn1a, Scnn1b and Scnn1c respectively [34]. Expression and activity of
ENaC, which represents the rate-limiting step for luminal Na+ entry, is under complex hormonal regulation, by aldosterone, vasopressin and insulin [35]. As a result of the failure to initiate cAMP-mediated chloride secretion, ENaC is activated, which increases sodium and water reabsorption in airway epithelia [27,
36, 37]. Furthermore, Legris et. al report that patients with CF exhibit a chronically driven renin-angiotensin-aldosterone axis to assist in the maintenance of extracellular fluid volume and compensate for salt-deficiency, which may contribute to ENaC increased expression and activity observed in human CF nasal epithelia [38, 39]. Regulation of ENaC by CFTR has been proposed as a
4 requirement for maintaining ASL height, which is critical for efficient mucociliary transport and removal of pathogenic bacteria [25, 36].
The low volume hypothesis correlates the inability of CF epithelia to transport chloride with disinhibition of ENaC. In support of this hypothesis, increased transepithelial sodium absorption in the airways of Scnn1b transgenic mice led to CF-like airway disease [27, 36]. These mice exhibit decreased ASL height, increased mucus concentration and adherence, neutrophilic inflammation and poor bacterial clearance similar to human CF airways [36]. Work from the laboratories of Boucher and Pilewski suggests that ENaC may be activated in CF airway epithelia as the result of an imbalance between serine proteases and protease inhibitors present in the airway surface liquid [27, 40]. Protease inhibitors present in normal ASL prevent ENaC activation by channel activating proteases, which cleave the α and γ subunits to activate ENaC. While neither group measured the concentration of proteases present in the ASL of cultures of
CF airway epithelial cells, they clearly demonstrated that ASL volume is important for modulating ENaC activity and that the normally silent ENaC is constitutively active in CF epithelia [40]. Using primary human tracheal epithelial cells, they also showed that addition of nystatin, a sodium ionophore, decreased
ASL height [41]. This effect, which was abrogated by amiloride, was recapitulated with bumetanide treatment to block chloride secretion and confirmed that coordinated sodium absorption and chloride secretion are required to maintain proper ASL height. Taken together, these studies provided the first clear mechanistic link between accelerated ENaC-mediated sodium absorption and
5 disease pathogenesis in the airway, highlighting the contribution of ENaC in vivo to ASL dehydration in the presence of functional CFTR.
The contribution of the Na,K-ATPase
The early appreciation that CF airway epithelia absorbed sodium at an accelerated rate led to the observation that nasal tissue from CF patients had a
60% increase in ouabain binding sites compared to non-CF tissue, indicating an increase in membrane localized Na,K-ATPase [42]. The Na,K-ATPase, identified in 1957, is a heteromeric enzyme composed of a large (∼100 kDa) catalytic α- subunit, a smaller (∼45 kDa) glycosylated β-subunit, and is regulated by
members of the FXYD protein family [43-45]. Four α isoforms (α1, α2, α3, α4),
three β isoforms (β1, β2, β3) and seven FXYD subunits (FXYD1-7) have been identified and are expressed in a tissue-specific manner [45-47]. An integral membrane protein, the Na,K-ATPase maintains the electrochemical sodium and potassium gradients across the plasma membrane by using the energy of ATP hydrolysis to exchange 3 intracellular sodium ions for 2 extracellular potassium ions [44, 48-50]. In airway epithelia this supports a transepithelial sodium axis, with sodium entering primarily through ENaC in the apical membrane and exiting through the basolateral Na,K-ATPase.
Airway epithelia express the α1 and β1 isoforms, which are coordinately increased in abundance around the time of birth, presumably to aid in lung liquid clearance [51-53]. Increased expression and activity of the Na,K-ATPase create a transepithelial osmotic gradient that causes water to move out of the airspace
6 into the interstitium and capillaries, which helps maintain appropriate ASL height
[54-57]. Similar to ENaC, Na,K-ATPase function in airway epithelia can be pharmacologically upregulated using β-adrenergic agonists, dopamine, aldosterone and epidermal growth factor to modulate liquid clearance in normal and injured lungs [58-60]. Overexpression of Na,K-ATPase subunits in vitro and in vivo have demonstrated that active sodium transport is critical for edema clearance and suggest that gene therapy delivering Na,K-ATPase subunits may protect the lung after injury [57, 58, 61].
The vectorial transport of sodium and water require that the airway epithelium maintain a polarized phenotype, and it has been shown that Na,K-
ATPase function has a role in the formation of junctional complexes between epithelial cells [62-65]. Inhibition of Na,K-ATPase activity with ouabain or by K+ depletion prevented tight junction formation and cell polarization [64, 66].
Expression of the heavily glycosylated β1 subunit is a key factor regulating cell- cell adhesion, which is independent of adherens or tight junctions [67]. Repletion of the β1 subunit in MDCK cells transformed with the Maloney-Sarcoma virus, which have decreased E-cadherin and β-subunit levels, restored cell polarization, decreased cell motility and increased adherence [66]. These observations suggest that disruption of cellular ion homeostasis maintained by the Na,K-
ATPase results in the generation of signals that regulate cell adherence, leading to the possible loss of cell structure and epithelial – mesenchymal transition.
The importance of the Na,K-ATPase to regulate epithelial cell polarization and motility is emphasized by observation that decreased alpha- and beta-
7 subunit expression in renal clear cell carcinoma correlates with higher grade tumors and metastatic potential [68, 69]. Similarly, expression of the α1 subunit was decreased in advanced prostatic carcinoma when compared to normal and benign prostatic hyperplastic tissue [70]. Prolonged exposure to ouabain detaches cells from their substrate, indicating the existence of a link between pump function and attachment [71, 72]. It is currently unclear whether diminished membrane localized Na,K-ATPase or decreased ion transport activity is responsible for these observations, but taken together imply that altering either the pump or attachment function of the Na,K-ATPase may lead to neoplastic development in epithelia. Furthermore, the role of the FXYD family proteins, which associate with the Na,K-ATPase and modulate its activity, have recently received attention as possible indicators of metastatic potential in addition to their effect on Na,K-ATPase pump kinetics [73-76].
The FXYD family: modulators of Na,K-ATPase activity
In 2000, Sweadner and Rael first defined the FXYD family, a small family of type-I proteins characterized by a 35-residue sequence containing an invariant proline-phenylalanine-X-tyrosine-aspartate (PFXYD) domain, two glycine residues in the single transmembrane domain, and a conserved intracellular serine [45]. Typified by the γ-subunit (FXYD2) of the renal Na,K-ATPase, the family also includes phospholemman (FXYD1), which regulates ion homeostasis in the cardiac sarcolemma; MAT-8 (FXYD3), a mammary tumor marker; CHIF
(FXYD4), or channel-inducing factor; RIC (FXYD5), for related to ion channel;
8 phosphohippolin (FXYD6), found primarily in the hippocampus and cerebellum; and FXYD7, which is primarily uncharacterized but found in the peripheral nervous system [73, 76-84]. Characterization of EST replicates with known proteins initially suggested that FXYD proteins did not share similar function, however work from several laboratories has confirmed that FXYD proteins interact with and modulate Na,K-ATPase kinetics [85, 86]. While not an integral part of the α/β complex, FXYD proteins are expressed in a tissue-specific manner and are believed to fine-tune Na,K-ATPase function according to the specific needs of particular tissue or physiological state [86, 87].
In addition to the various combinations of α and β isoforms, FXYD proteins represent another mechanism for controlling enzyme activity. The main function of the Na,K-ATPase is to maintain low intracellular sodium concentrations, which can be regulated via hormones for rapid (short-term) or sustained (long-term) effect. FXYD proteins exert their effects by interacting with the Na,K-ATPase and modulating the maximum catalytic activity, enzyme substrate affinity for Na+ and/or K+, and on apparent ATP affinity [87]. Ectopic expression in Xenopus oocytes and mammalian cells has revealed that the transmembrane domain mediates the effects of FXYD proteins on Na,K-ATPase cation affinities and that residues within the FXYD extracellular domain affect ATP affinity [88-91].
Immunofluorescence analysis has shown that FXYD1-4 and FXYD7 colocalize with the Na,K-ATPase in the basolateral membrane, and coimmunoprecipitation studies have demonstrated α/β/FXYD complex formation in solublized membrane preparations [92-95]. The demonstration of these associations has confirmed that
9 FXYD proteins act as auxiliary subunits of the Na,K-ATPase and modulate enzyme activity, with each family member possessing distinct regulatory effects.
The main rate-limiting step in Na,K-ATPase activity is the deocclusion of
+ cytoplasmic K , or the E2(K)-ATP to E1-ATP transition, induced by low-affinity
ATP and Na+ binding (Figure 1-1) [86, 87, 96]. Evidence that FXYD proteins affect the kinetics Na,K-ATPase activation was obtained using an antibody raised against a C-terminal epitope of FXYD2 (γ subunit), which partially inhibited the activity of purified renal Na,K-ATPase [83, 97, 98]. Later, overexpression of
FXYD2 isoforms in HEK 293 cells was found to reduce the Km for activation by
ATP in microsome preparations, and subsequent studies in the rat renal cell line
NRK-53E have demonstrated that the γa subunit (a splice variant of FXYD2) modulates both Na+ and K+ affinities [98, 99]. While experimental systems vary in their estimation of the modulatory effects of FXYD2 on the K1/2 for extracellular
+ + K , it is clear that γ raises the K1/2 for cytoplasmic Na approximately 2-fold [86].
In contrast, FXYD4 (CHIF) induces a two- to threefold decrease in the K1/2 for intracellular Na+ and has little affect on the K+ activation [78, 95, 96, 100]. Studies in FXYD4 knockout mice, which demonstrate decreased K+ excretion under low
Na+ intake, confirm that FXYD4 activates renal Na,K-ATPase by increasing the apparent affinity for Na+ [101, 102]. The observation that FXYD2 and FXYD4 have opposite functional effects, consistent with their different patterns of expression along the nephron, were pivotal in recognizing that FXYD proteins might be tissue specific regulators of Na,K-ATPase activity. Currently, FXYD1-4 and FXYD7 have been shown to associate with and modulate the kinetic
10 properties of the Na,K-ATPase in a tissue-specific manner. As a result of the initial study indicating that FXYD5 was highly expressed in transport tissues, such as the lung, we hypothesized that FXYD5 was a regulator of Na,K-ATPase activity in airway epithelial cells.
FXYD5
For many years, the γ-subunit of the Na,K-ATPase was an orphan glycoprotein, remarkable only for its association with the sodium pump, and little was known of its function. Although the classification of the FXYD protein family in 2000 identified new siblings, the function of most of these proteins and their ability to affect ion transport remained unknown [45]. FXYD5, also known as
Related to Ion Channel (RIC), was originally identified as a gene upregulated in fibroblasts transformed with the E2a-Pbx oncoprotein, which transforms T lymphoblasts and blocks myeloblast differentiation in mice (Figure 1-2) [103].
This was the first study to demonstrate that FXYD5 was normally expressed in the lung, spleen, skeletal muscle and testes. FXYD5 also demonstrated biphasic expression during development, similar to observations that other FXYD proteins are required for blastocoel cavitation. This observation allowed speculation that
FXYD5 expression in the lung may be important for post-natal clearance of lung liquid [104]. In 2002, FXYD5 was identified as a 45 kDa glycoprotein by the
Hirohashi group in an immunological screen for cancer-associated cell membrane glycoproteins [105]. They found that overexpression was associated with proportionate downregulation of E-cadherin in a liver-cancer cell line,
11 resulting in decreased cell-cell adherence. An increased number of metastatic nodules was observed after injection of FXYD5-overexpressing cells into immunodeficient mice compared to the same mock-transfected cell line. Based on their observations using human FXYD5 in their overexpression experiments, they originally named FXYD5 “Dysadherin” ostensibly due its properties as an anti-adhesion molecule [105].
Soon after, the same group correlated increased FXYD5 expression with tumor aggressiveness in pancreatic ductal adenocarcinoma. Importantly, they found that FXYD5 was expressed at the cell membrane of cancer cells, but not in nontumor or tumor-adjacent cells, suggesting that altered expression of FXYD5 may be an indicator of metastatic potential. Though they did not demonstrate whether FXYD5 upregulation was the result or cause of increased tumor metastases, they showed that FXYD5 expression was stronger in infiltrative and poorly differentiated tumor nests [106]. These observations were further substantiated with the observation that siRNA directed against human FXYD5 altered cell morphology and decreased cell motility in a pancreatic cell line. Cells transfected with FXYD5 siRNA have a more spread shape, are larger and have increased transverse actin stress fibers commonly associated with a non-motile phenotype when compared to control cells [107]. As a result of these studies, the
Hirohashi group evaluated FXYD5 expression in thyroid, pancreatic, esophageal, colon, prostate, breast, tongue, liver, testicular, head/neck, non-small cell lung and stomach cancers and found that increased FXYD5 expression was significantly associated with poor patient prognosis, and increased membrane-
12 localized FXYD5 indicated tumor aggressiveness [105-115]. It is thought that
FXYD5 exerts these effects by altering the cell-adhesion system, particularly by down-regulating E-cadherin expression, but the molecular mechanisms responsible have not been elucidated.
Studies have shown that epithelial cell-cell adhesion is dependent on E- cadherin mediated regulation of tight junctions, the loss of which may result in the transition to a mesenchymal phenotype and lead to carcinoma development.
The data suggest a relationship exists between FXYD5 and E-cadherin expression, but it is unclear whether expression is inversely related or independent [116]. Knockdown of FXYD5 in an siRNA treated breast cancer cell line identified chemokine (C-C motif) ligand 2 (CCL2) as the transcript most affected by FXYD5 downregulation [117]. Further analysis implied that the ability of FXYD5 to promote cell motility and metastasis was dependent on the establishment of a CCL2 autocrine loop, but it is possible that this was due to undetermined effects resulting from increased exposure to CCL2 [116]. Similarly,
Na,K-ATPase function is critical in the establishment of epithelial cell polarity and adherence. Disruption of E-cadherin function and decreased expression of Na,K-
ATPase subunits has also led to alterations in the rate of epithelial wound healing, which requires cells to mobilize and migrate into a wound site [118].
While the Hirohashi laboratory has demonstrated that FXYD5 may regulate cell motility and is increased in many carcinomas, only recently has evidence been provided that demonstrates FXYD5 may interact with and modulate Na,K-
ATPase function.
13 Work from the laboratories of Karlish and Garty, two pioneers of the FXYD field, has recently focused on murine FXYD5, which they report to be expressed as a 25 kDa protein in the lung, heart, kidney, spleen and intestine [119].
Previous reports have shown that FXYD2 (γ) and FXYD4 (CHIF) are specifically expressed in the proximal convoluted tubules or distal convoluted tubles of the kidney, respectively. However, Lubarski et. al. demonstrate that FXYD5 is also expressed in the kidney, though only in the collecting ducts [119]. These data support the notion that each FXYD protein is expressed in a tissue-specific manner, but suggest further regulation may be implemented at the cellular level or in response to a physiological stimulus. In particular, although FXYD5 has been documented from multiple whole-organ preparations, FXYD5 expression may be dependent on unknown cell-type specific factors.
While altered FXYD5 expression has been associated with perturbations in cell adhesion, few studies have evaluated whether FXYD5 affects Na,K-
ATPase function. New work has now shown that FXYD5 co-immunoprecipitates with the α1 subunit of kidney Na,K-ATPase and induces a two-fold increase in ouabain-inhibitable current in Xenopus ooctyes injected with murine FXYD5 cRNA compared to control-injected ooctyes [119]. Measurements on the K+ current activation curve indicate that FXYD5 may increase Vmax and the K0.5 for external K+ [119]. Similarly, 86Rb+ uptake was increased almost two-fold in
Xenopus ooctyes. However, these authors also overexpressed the rat α1 and pig
β1 Na,K-ATPase subunits in their experimental system, which have different affinities for Na+ and K+ and undefined interactions with FXYD proteins. This may
14 account for the differences observed in Na,K-ATPase subunit expression in these studies [119, 120].
Collectively, the data indicate that a) FXYD5 is upregulated in numerous tumors; b) increased expression is a prognostic indicator of metastatic potential; c) overexpression correlates with increased motility, whereas knockdown increases cell adherence; d) associates with the Na,K-ATPase; and e) interaction with the Na,K-ATPase modulates pump function. It is currently unclear how interaction with the Na,K-ATPase modulates pump function and whether this is affected in epithelia where ion transport homeostasis is disturbed. Given the chloride transport abnormality in CF, the existence of CF models with and without consistent concomitant abnormalities in ENaC function, and the existence of
ENaC overexpressing mice, CF airway epithelia may therefore present an attractive model to help elucidate how FXYD5 modulates the kinetics of Na,K-
ATPase activation. Exploring this model may also explain how FXYD5 expression mediates epithelial cell motility, providing insight for future therapies directed towards anti-cancer drugs.
15
Forward ADP Na
Na E1-P Na Na Na Na - E1-ATP Na - E2-P
Na Na
3 Na+ 3 Na+ E2-P E1-ATP + 2 K 2 K+
K K - E1-ATP - E2-P K K
K K E2-ATP E2-P K K
ATP Pi
Intracellular Extracellular
Figure 1-1. Modified Post-Albers scheme. Adapted from Pederson et. al and
Holmgren et. al, showing the translocation of Na+ and K+ through the Na,K-
ATPase [121, 122]. E1 phosphorylation by ATP occludes three cytoplasmic Na+ eliciting a change in enzyme conformation to E2. The three bound Na+ ions are exported to the extracellular medium, which induces the binding of two K+ and enzyme dephosphorylation. ATP binding shifts the enzyme back to the E1 conformation, which elicits K+ import to the cytoplasm and Na+ binding to complete the cycle.
16
P F F Y D
Figure 1-2. Human FXYD5 cDNA and protein sequence. FXYD5 is a type-1 transmembrane protein, contains 178 amino acids, and is identified by a characteristic PFXYD domain (bold) and single transmembrane domain (boxed).
17 Chapter 2: Characterization of human and mouse FXYD5
Timothy J. Miller1,2 and Pamela B. Davis2 From the Departments of 1Pharmacology and 2Pediatrics, School of Medicine, Case Western Reserve University, Cleveland, Ohio
Abstract
FXYD5, also known as Dysadherin, was originally identified as a gene induced in NIH-3T3 cells transformed with the E2a-Pbx1 oncogene and subsequently cloned and characterized as a cancer-associated cell membrane glycoprotein. While both human and mouse FXYD5 encode a 178 amino acid protein that includes a putative signal sequence, a single transmembrane domain and a short cytoplasmic tail, the literature reports significant differences appear regarding each species molecular weight, migration and tissue expression. RT-
PCR analysis indicated that FXYD5 is expressed in Calu-3, HEK293, A549,
MEF, MDCK, LA4, NIH3T3, HuH7 and the previously characterized negative
PLC/PRF5 cell line. We compared Unigene expression profiles to determine the tissue specificity of human and mouse FXYD. Northern blot and quantitative RT-
PCR analysis confirmed that FXYD5 was highly expressed in murine lung tissue.
We developed a FXYD5 antibody (562), raised against the murine C-terminus, which was specific for murine FXYD5. Immunohistochemistry demonstrated specific staining for FXYD5 in the basal layer of murine epidermal tissue.
Immunoblot analysis demonstrated that murine FXYD5 was expressed in the liver, lung, kidney, spleen and pancreas. To better evaluate differences between human and mouse FXYD5, Flag-tagged FXYD5 chimeras were constructed which revealed that mature, fully glycosylated human FXYD5 migrated at
approximately 37 kDa, whereas mouse FXYD5 migrated at 25 kDa. These results document that FXYD5 is similarly expressed in human and mouse tissues, but that each ortholog is fundamentally different.
Introduction
The advent of high-throughput gene expression profiling has increased our ability to compare patterns of human and mouse gene expression over the past 30 years [123]. Mouse models, often used to study human genes, are believed to recapitulate human phenotypes due to the similar expression and function of orthologous genes [124]. While these direct evolutionary counterparts or gene pairs may be expected to retain the same function, the expression of each particular gene is often neglected within the context of the organism in order to examine that function [125]. As a result, expression profiles of human- mouse orthologs may be more divergent than expected, causing correlations drawn between supposedly identical proteins to be erroneous.
Such a difference is observed between the rat and human α1 subunit of the Na,K-ATPase [126, 127]. The 110 kDa catalytic α1 subunit is ubiquitously expressed, however the rat isoform is ten-fold more resistant to the cardiotonic steroid ouabain compared to the α1 subunit isolated from human, mouse, pig or dog [126, 128-130]. This difference has been exploited to develop model systems useful for the kinetic analysis of human Na,K-ATPase subunits, such as the newly identified FXYD family [78, 95, 98, 131-133].
19 FXYD5, also known as dysadherin and Related-to-Ion Channel (RIC), has been identified as a tumor-associated cell membrane glycoprotein in human cancer cells [105]. FXYD5, which is unique in the FXYD family as a result of an extended extracellular domain, appears to be highly expressed in the basal layer of squamous epithelia, endothelia and lymphocytes. Expression is particularly high in the epithelia of ion transport tissues, such as the lung. Initial reports indicate that overexpression of human FXYD5 downregulates E-cadherin in cancer cells, suggesting that FXYD5 may modulate epithelial cell-cell adhesion in a manner opposite that of the Na,K-ATPase β-subunit [66, 68, 134]. Sequence analyses of human and murine FXYD5 indicate similar full-length proteins, but recent reports suggest a disparity exists between predicted and reported size
[119].
Similar to other FXYD proteins and Na,K-ATPase subunits, FXYD5 may exist as multiple isoforms to allow for differential regulation of the Na,K-ATPase at various levels (i.e – transcriptional or translational) [120, 130]. Biochemical data suggest that human FXYD5 is approximately 45 kDa, whereas mouse
FXYD5 is 25 kDa. Neither protein contains Asn-X-Ser/Thr tripeptide sequences known for potential N-glycosylation, however both have extracellular domains that are rich in Ser-, Thr-, and Pro-rich residues, which can allow O-glycosylation
[135]. Identifying species specific differences may be a valuable tool for evaluating the effectiveness of FXYD5 model systems. In this chapter, we show that human and mouse FXYD5 share similar expression patterns but are fundamentally different proteins.
20 Methods and materials
Unigene profiles. Tissue expression profiles were generated using Homo sapiens sequence tag Hs.333418 and Mus musculus sequence tag Mm.1870 from the
National Center for Biotechnology Information (NCBI) Unigene profile at http://www.ncbi.nlm.nih.gov/sites/entrez?db=unigene&cmd=search&term=fxyd5.
Biochemical analysis and DNA manipulation. Human and mouse FXYD5 protein and DNA constructs were created using Genbank accession numbers
NM_014164, human and NM_008761, mouse and analyzed using Vector Nti
(Stratagene, Inc., Cedar Creek, Texas).
Cell lines. The mouse lung epithelial cell line LA4, human embryonic kidney
(HEK293), Madin-Darby canine kidney (MDCK), human lung adenocarcinoma
(Calu-3), mouse fibroblast NIH 3T3, mouse embryonic fibroblast (MEF), human lung carcinoma (A549), human hepatoma (HuH7) and human liver hepatoma
(PLC/PRF5) cell lines were obtained from the American Type Culture Collection
(Manassas, VA). HEK293 and Calu-3 cells were cultured in Earles Minimum
Essential Medium (EMEM; Cellgro, Mediatech, Inc., Herndon, VA), A549 and
LA4 cells were grown in Kaighn’s modification of F12 medium (F12K, Cellgro),
NIH3T3 and MEF cells were grown in Dulbecco’s Modified Eagle medium
(DMEM, Cellgro), PLC/PRF5 and MDCK cells were grown in Eagle’s minimum essential medium (Cellgro). All media were supplemented with 10% heat- inactivated FBS and cells were grown in a 37°C, 5% CO2-95% 02 atmosphere.
21
Mouse strains. Animals were maintained in the CF Animal Core Facility at CWRU before use and all studies were performed under CWRU IACUC approved protocols. Breeding pairs of heterozygote congenic mice (>N10) bearing the
S489X mutation (B6.129P2-Cftrtm1unc, stock no. 2196) and C57BL/6J mice were purchased from Jackson Laboratory (Bar Harbor, ME). Breeding pairs of heterozygote mice bearing the ΔF508 Cftr mutation in mixed genetic background were a kind gift from Dr. Kirk Thomas from the University of Utah, and were backcrossed into the C57BL/6J background for at least 10 generations in the CF
Animal Core Facility at CWRU before use. Cystic fibrosis mice for these strains are indicated by their Cftr mutation and are referred to as “CF mice”.
RNA isolation. Total RNA was isolated from approximately 30 mg tissue from 4 to
6 week old C57BL/6J mice or from 10 cm dishes of LA4 or HEK293 cells using the RNAprotect kit according to the manufacturers instructions (Qiagen, Inc.) and stored at –80ºC. RNA was quantified on a spectrophotometer and visualized by agarose gel electrophoresis to determine quality.
RT-PCR, cloning and site-directed mutagenesis of FXYD5. The Superscript II
One-Step RT-PCR kit (Invitrogen, Inc., Carlsbad, CA) was used to reverse transcribe and amplify FXYD5 from LA4 and HEK293 cells. FXYD5 cDNA was isolated by RT-PCR using primers designed from accession numbers NM014164
(human) and NM008761 (mouse), which contained a HindIII and NotI restriction
22 site on the 5’ and 3’ end respectively. The following primers were used to generate FXYD5 clones: Human 5’AAGCTTGCTAGCGCCG
CCACCATGTCGCCCTCTGGTCGCCTGTGTCT (forward), 5’ AGTCGTCTA
GATCACCTGCAACGATTCCGGCATAAC (reverse); mouse 5’ AAGCTTGCT
AGCGCCGCCACCATGTCACTGTCCAGTCGCCTGTGTCT (forward) 5’
AGTCGTCTAGATCACCTGTG GCGATTCAGGCAAATT (reverse). The following
RT-PCR conditions were used: reactions were incubated at 50ºC for 30 minutes, followed by 2 minute initial denaturation at 95ºC and 40 cycles of 94ºC, 1 minute denaturation, 1 minute 58ºC primer annealing, 45 seconds of primer extension.
RT-PCR products were digested with HindIII and NotI restriction enzymes and agarose gel purified using the Qiaquick gel purification kit (Qiagen,Inc., Valencia,
CA). cDNAs were subcloned in pBSK2 vector to create pBhF5k (human) and pBmF5k (murine) (Figure 2-8). To generate an N-terminus Flag tag in human
FXYD5 that did not alter the N-terminus signal sequence, codon Q22 was mutated using the Quickchange site-directed mutagenesis kit (Stratagene, Inc.,
Cedar Creek, Texas). A silent mutation was introduced (CAG mutated to CAA) to create a new restriction site, Acl1. The following sequence was then inserted in frame at the Acl1 site to produce pBhF5kQ22Flag: 5’
CGGATTACAAAGATGATGATGATAAGA 3’. To create a C-terminal Flag-tagged
FXYD5, pBhF5k was digested with NotI and TfiI and agarose gel purified. The following oligo was then ligated to produce pBhF5kFlag: 5’
AATCGTTGCAGGGATTACAAAGATGATGATGATAAGT GATCTAGAGC.
Similarly, a C-terminus Flag-tag was inserted into murine FXYD5 to create
23 pBmF5kFlag. The original and modified versions of FXYD5 were then subcloned into the previously described pKCERegfpSV expression vector (31) using
HindIII/Not to create pKCERhF5k, pKCERmF5k, pKCERhF5kQ22Flag, pKCERhF5kFlag and pKCERmF5kFlag. Human and mouse chimeric constructs were created by utilizing the AlwnI restriction site to produce pkCERhF5kQ22FlagmF5, pKCERhF5kmF5Flag, pKCERmF5khF5Flag (Figure 2-
8). A truncated version of murine FXYD5 was created by digesting pKCERmF5kFlag with AlwnI and HindIII, filling in both 5’ overhands with Klenow and re-ligating the vector to make pKCERmF5kFlagtrunc. All vectors were sequenced on both strands to validate RT-PCR and cloning procedures to ensure cDNA open reading frames were in-frame.
Northern blot analysis of FXYD5 tissue expression. 5 µg of total murine RNA was separated on a 1% agarose/formaldehyde gel at 100V for 3 hours and transferred overnight in 10x SSC via a Turboblotter (Stratagene, Inc., Cedar
Creek, Texas) onto nytran membrane (Invitrogen Inc., Carlsbad, CA). The nytran membrane was then fixed in 5% glacial acetic acid, washed in water, and stained with 0.04% methylene blue to assess transfer and identify 28S and 18S rRNA.
The nytran membrane was then prehybridized for 20 minutes at 68ºC in
Quickhyb solution (Stratagene, Inc., Cedar Creek, Texas, Inc.). 30 ng of murine
FXYD5 cDNA was prepared using the Multiprime labeling kit (Amersham, Inc.) with 50 µCi α-P32-dCTP, and the blot was hybridized for 1 hour at 68ºC. The blot
24 was then washed two times 10 minutes each in 2x SSC, 0.1% at room temperature and in 0.2x SSC, 0.1% SDS and exposed for 36 hours on film.
Quantitative RT-PCR. 0.5 µg of total RNA was utilized to synthesize cDNA with the First Strand cDNA synthesis kit (Roche Molecular Biochemicals, Mannheim,
Germany, Inc.). cDNA synthesis was performed using oligo-p(dT)15 primer per kit instructions and stored at -20ºC. Quantitative PCR was performed using the
Lightcycler FastStart DNA master Sybr green kit (Roche Molecular Biochemicals,
Mannheim, Germany, Inc.). The primer sequences used for human and mouse
FXYD5 are: human 5’ ATGTCGCCCTCTGGTCGCCTG (forward), 5’
TCACCTGCAACGAT TCCGGCA (reverse) and mouse 5’
ATGTCACTGTCCAGTCGCCTGTGT CTCCTCACT (forward), 5’
TCACCTGTGGCGATTCAGGCAAAATTGAGACAA (reverse). DNA was amplified using a Roche Lightcycler with the following parameters: 95ºC 10 minute initial denaturation followed by 35 cycles of 95ºC 10 second denaturation,
58ºC 7 second anneal, 72ºC 10 second extension. Copy number was measured against a standard curve of linearized plasmid DNA and normalized to GAPDH.
Second point derivatives were used for PCR analysis. Specificity of DNA amplification was assessed by melting point analysis and confirmed by agarose gel electrophoresis.
Preparation of polyclonal FXYD5 antibody. The sequence
GKCRQLSQFCLNRHR was used by Covance, Inc. (Princeton, NJ) to create rabbit polyclonal antibodies directed against FXYD5. Antisera were screened by
25 ELISA and analyzed by immunoblot to identify successful immunization. Serum was collected, stored at -20ºC and affinity purified. FXYD5 562 polyclonal antisera recognized mouse FXYD5 as analyzed by immunohistochemistry of murine squamous epithelia and immunoblot analysis (Figure 2-6).
Isolation of crude membranes. Mouse tissue or cultured cells were isolated, washed with 25 mM imidazole, 1 mM EDTA, 250 mM sucrose and protease inhibitor cocktail (Sigma, St. Louis, MO) and manually homogenized with 25 strokes of a prechilled Dounce homogenizer. Nuclei, unbroken cells and mitochondria were separated by centrifuging at 6000 x g for 5 minutes. The supernatant was collected and saved, while the pellet was homogenized again and centrifuged. The two supernatants were combined and centrifuged at
125,000 x g to pellet crude microsomal membranes. The pellets were resuspended in 25 mM imidazole, 1 mM EDTA, 20 mM NaCl and stored at 4ºC.
Immunoblot analysis of FXYD5. Crude membranes (10-20 µg) were dissolved in
30 µl 2x Laemmli buffer, incubated at 37ºC for 20 minutes, loaded onto 12%
SDS-PAGE and resolved at 100 V for 1.5 hr. Proteins were transferred via semi- dry blotting onto nitrocellulose (Bio-Rad, Hercules, CA), blocked with 1% BSA and incubated with either rabbit polyclonal anti-FXYD5 562 (1:250) antibody or antibody against the M2 Flag tag (1 mg/ml) (Sigma, St. Louis, MO) in 1%
BSA/Tris-buffered saline for 1 hr at room temperature. The blot was then
26 incubated with species specific secondary antibodies (1:4000) conjugated to
HRP and visualized using Pierce’s Supersignal Pico West detection kit.
Immunohistochemistry of mouse tissue. Adult homozygous ΔF508 (CF) mice and their wild-type littermates were sacrificed and fixed by perfusion using 2% paraformaldehyde in 1x PBS (PH 7.4) via left ventricle. Lung tissue was immersed in the same fixative solution for 48 hrs at room temperature, dehydrated and embedded in paraffin using standard histological procedures. 5
µm sections were made and mounted onto the slides with Fisherbrand superfrost plus coated with Vectabond (Fisher 12-550-15). After deparaffinization, the sections were treated with cold methanol for 15 minutes followed by 1% hydrogen peroxide in distilled water for 30 minutes at room temperature to block endogenous peroxidase activity, washed with PBS and incubated with 1.5% normal goat serum/PBS (Vector Laboratories, PK-6101) for 4 hrs at room temperature. The sections were then incubated with 6.8 µg/ml of affinity purified
562 polyclonal rabbit antibody against FXYD5 overnight at 4ºC. Blocking solution without primary antibody was used as a negative control. After washing 3 times in 1x PBS, the sections were incubated with 1.5% biotinylated goat anti-rabbit
IgG antibody 1 hr at room temperature. After washing 3 times with PBS, the sections were incubated with Vectastain ABC reagent for 30 min and washed again. The sections were developed with 3,3'-diaminobenzidine (DAB) using
DAB subtrate kit (Vector Laboratories, SK-4100) for 5 minutes. After washing 2x
27 with water, the sections were counterstained with hematoxylin (Themo Electron
Corporation, 6765004).
Indirect immunofluorescence. HEK293 cells were cultured in 4-well chamber slides (Permanox) to 80% confluence and transfected with pKCERhF5kQ22Flag with 4.5 µl Lipofectamine 2000 (Invitrogen, Carlsbad, CA) and 1.5 µg DNA per well. At 24-36 hrs post-transfection, monolayers were washed once with phosphate buffered saline (PBS), fixed with 2% paraformaldehyde/PBS for 10 minutes and washed with PBS twice. Non-specific antibody binding was blocked by addition of 4% bovine serum albumin (Gibco) for 45 minutes. Primary anti-M2
Flag mouse antibody was diluted in 1% BSA/PBS at 2 mg/ml, incubated on monolayers for 45 minutes and aspirated. Monolayers were washed twice with
PBS and then incubated in 1%BSA/PBS containing goat anti-mouse IgG Alexa-
Fluor 568 (1:250, Stratagene, Inc., Cedar Creek, Texas) for 45 minutes. Cells were then washed twice, the nuclei counterstained with Hoechst (Stratagene,
Inc., Cedar Creek, Texas) and mounted with Fluormount-G. Slides were allowed to dry overnight and immunofluorescent localization of FXYD5-Flag was assessed on a Zeiss 200M Axiovert inverted microscope, with a DG4 switchable fluorescent light source (Sutter Instrument Company, Novato, CA) and a 12-bit
CoolSnap HQ camera (Roper Scientific, Tucson, AZ) under control of MetaMorph v 6.2 (Molecular Devices, Sunnyvale, CA. Images were obtained with a 63X numerical aperture 1.3 fluar lens using excitation and emission filter passbands
28 of 260±20nm and 645±30nm, respectively. Typical exposure times for individual frames were 200 ms.
29 Results
Comparison of Human/Mouse FXYD5 tissue expression patterns
Our understanding of FXYD5 is based on the initial identification of human, mouse and rat cDNA and protein sequences, which share similar lengths and conserved residues. However, our biochemical comparison of human and mouse FXYD5 indicate significant differences may exist (Table 2-1). Mouse
Fxyd5, located on chromosome 7B1, is orthologically similar to human FXYD5, located on chromosome 19q12-q13.1. Sequence comparison shows that the mRNA for mouse FXYD5 is 1144 bp, compared to 890 bp for human FXYD5.
The amino acid sequences calculated from the molecular weights of the human and mouse FXYD5 cDNA sequences, which are 54.5% homologous, are 19.47 and 19.45 kDa, respectively. Although both full-length proteins are 178 a.a., the isoelectric point and charge at pH 7.0 are very different, which suggests that the species of origin may account for differences observed during immunoblot analysis. More importantly, these differences may determine the parameters of kinetic effects on Na,K-ATPase activity.
Unigene is essentially an automated analytical system for producing an organized view of the transcriptome of a particular species or tissue. We analyzed Unigene expression profiles for human and mouse FXYD5 to compare tissue expression patterns. The profile of human FXYD5 is composed of
6,808,401 sequences, including 12 mRNA and 378 ESTs, whereas mouse
FXYD5 has 4,204,718 sequences, including 8 mRNA and 185 ESTs (Figures 2-1 and 2-2). Digital Differential Display (DDD) analysis did not yield significant intra-
30 species tissue expression pattern differences as a result of the requirement of over 1000 sequences necessary for the Fisher Exact test. However, the human
Unigene profile confirmed the observations of prior studies, which reported that
FXYD5 is highly expressed in lymphocytes (lymph node), lung, blood, bone marrow, pancreas and skin, whereas a notable lack of expression was observed in the tissues of the CNS and reproductive systems. However, the human
Unigene profile indicates a previously unreported high level of expression in the salivary gland, stomach, tonsil and umbilical cord. While the murine tissue expression profile was similar to human, there were a few notable exceptions.
While Fxyd5 was similarly expressed in the skin and lymph, a large difference in relative transcript-per-million (TPM) expression was observed in the blood, pineal gland and uterus compared to the human profile. This was also observed when compared to expression in the stomach and pancreas. Taken together, these results suggest that similarities exist in tissue-specific expression patterns between species but that FXYD5 may have other, unknown species-specific roles in particular tissues.
The general pattern of human tissue expression suggested that FXYD5 was to a large extent ubiquitously expressed. To assess the validity of the murine
Unigene profile by more conventional methods, we assayed murine Fxyd5 expression by Northern blot analysis. We found that Fxyd5 was strongly expressed in preparations of total lung and spleen tissue, with a prominent signal also observed from the trachea (Figure 2-3). Quantitative RT-PCR analysis confirmed that total lung tissue had a 5-fold stronger signal compared to
31 intestine, liver, heart, brain and kidney after normalization to Gapdh expression
(Figure 2-4), suggesting that FXYD5, may be the lung-specific regulator of the
Na,K-ATPase in the mouse.
We examined FXYD5 expression in a panel of human and mouse cell lines to identify a FXYD5-negative control line for future studies. In contrast to prior reports, RT-PCR indicated that the human PLC/PRF5 hepatoma cell line was positive for the 556 base pair band indicating FXYD5 expression, as did all other cell lines tested (arrow, Figure 2-5). Interestingly, cell lines derived from human lung carcinomas, such as Calu-3 and A549 (Figure 2-5, lanes 2, 4), had the strongest expression of FXYD5, which was confirmed by quantitative RT-
PCR (data not shown). Smaller bands of 200 and 250 bp correspond to cDNA products resulting from internal start codons. Based on these results, we conclude that the Madin-Darby canine kidney cell line (MDCK) may be suitable for future studies analyzing the effects of FXYD5 on Na,K-ATPase ion transport kinetics.
We next developed a rabbit polyclonal antibody raised against a C- terminus epitope that recognizes mouse FXYD5 (Figure 2-6A). Our antibody recognized a 27 kDa protein in membrane preparations from murine airway LA4 cells, which was absent on immunoblots blocked with immunizing peptide or on immunoblots probed with control antisera (data not shown). We modified murine
FXYD5 to contain a C-terminus M2-Flag epitope tag, and immunoblot analysis confirmed that murine FXYD5-Flag was approximately 27 kDa (Figure 2-6B). A slight increase in electrophoretic mobility was observed, possibly due to the
32 increase in negative charge, however this phenomenon has previously been observed in other Flag-modified proteins. Prominent FXYD5 expression has been previously documented in epidermal tissue, and in situ immunohistochemistry with our FXYD5 antibody 562 comparably recognized
FXYD5 in murine squamous epithelia compared to sections stained with non- specific antibody (Figure 2-7A,B) [111, 113, 136]. The development of a FXYD5 antibody allowed us to determine whether Fxyd5 mRNA transcription correlated with FXYD5 expression in various murine tissues. Immunoblot analysis indicate that FXYD5 was strongly expressed in the liver, lung, kidney, spleen, and pancreas, suggesting that Fxyd5 may be differentially regulated or that the half- life of mature protein is different in each tissue (Figure 2-8). These results document that murine FXYD5 is approximately 27 kDa and confirm that human and mouse FXYD5 are not identical proteins.
Analysis of human and mouse FXYD5. Previous studies demonstrated species- specific, unexplained disparities between human and murine FXYD5. Although both human and murine FXYD5 have the same calculated molecular weight
(Table 2-1), human FXYD5 migrates at apparent MW 35-50 kDa and mouse
FXYD5, at 25 kDa [105, 119]. Human FXYD5 has a large stretch of negative charge in the C-terminal portion of the extracellular domain, but mouse FXYD5 does not, causing a large difference in overall molecular charge at pH 7.0 and very different isoelectric points (Figure 2-9). To test whether this domain accounts for the species-specific variation, we created chimeric constructs of N-
33 and C-terminal Flag-tagged fusion proteins (Figure 2-10), then transfected
HEK293 and LA4 epithelial cells. The species of origin of the C terminus determined the apparent molecular weight on SDS-PAGE whether the transfections were into mouse LA4 cells or human HEK293 cells, (data not shown), demonstrating that the apparent molecular weight differences are not due to the cell model. Comparison of an N-terminus Flag-tagged human FXYD5
(Figure 2-10B, lane 3) to C-terminus Flag-tagged (Figure 2-10B, lane 2) shows that placement of the Flag tag does not significantly alter the apparent MW of human FXYD5. We exchanged the N- or C-terminus halves of human and mouse
FXYD5 using the AlwnI site. No shift in apparent MW was observed in human
FXYD5 when the human N-terminus was replaced with mouse sequence
(compare Figure 2-10B lanes 3 and 5). However, when the human C-terminus was replaced with mouse sequence a decrease in the apparent MW was observed (compare Figure 2-10B lane 3 with lanes 4 and 6). Conversely, when the N-terminus of mouse FXYD5 was replaced with the corresponding human sequence, only a small increase in apparent MW was observed (compare Figure
2-10B lane 9 with lanes 10 and 12). A much larger increase in apparent MW occurred when the mouse C-terminus was replaced with human sequence
(compare Figure 2-10B lane 9 with lane 11). A truncated version of mouse
FXYD5-Flag was created (Figure 2-10B, lane 8) to compare the unprocessed
MW of FXYD5 and the use of alternative start codons (Figure 2-10B arrows).
Although location of the Flag tag did not affect the apparent molecular weight of either species, the C-terminus Flag tag is inefficient for
34 immunofluorescent detection in vivo (data not shown), as others have shown
[119]. The Q22Flag construct, however, is efficiently detected in the membrane of HEK293 cells (Figure 2-10C, D). Similar results were observed in LA4 cells
(data not shown). In HEK 293 cells transiently transfected with the pKCERhF5kQ22Flag, there is clearly a stronger localization of FXYD5 at homotypic cell borders suggesting that FXYD5 may have a role in cell-cell adhesion, although the mechanism governing this interaction is unknown [75].
35 Discussion
Initially, Fxyd5 was shown to be highly transcribed in mouse spleen and lung, with moderate expression in the heart, kidney and skeletal muscle [103]. A subsequent study showed that FXYD5 was strongly expressed in the lung, spleen, kidney, and intestine, with little or no expression in the liver, heart and skeletal muscle. An alternatively spliced isoform was also identified in the brain
[119]. The same group later published data demonstrating that FXYD5 was largely expressed in the lung, liver, and spleen, with moderate expression in the kidney and heart and no expression in the brain or skeletal muscle [120]. The data collectively suggest a high amount of FXYD5 expression in the lung, spleen and kidney, however a disparity exists whether FXYD5 is expressed in the brain, intestine and skeletal muscle.
Establishment of the dbEST database in 1992 was invaluable for the discovery of new genes and served as a collection point for new ESTs [123,
137]. EST registration comprises a large amount of Genbank, accounting for nearly two-thirds of all new submissions [137]. Although ESTs may be an inaccurate indicator of gene expression, the total number of ESTs and the tissue source of the libraries of origin may be useful for interspecies comparison [137].
In this study, we utilized the Unigene database to compare tissue expression profiles of human and mouse FXYD5. Furthermore, we biochemically compared human and mouse FXYD5 in an effort to clarify apparently discrepant data presented in previous reports.
36 A comparison of Unigene tissue expression profiles confirmed that human and mouse FXYD5 is highly expressed in the skin, blood, lung and spleen.
However, there are many cell types present in whole tissue preparations, and we acknowledge that caution should be used when interpreting profiles from whole- tissue preparations. It is interesting to note specific differences exist between certain tissues; human FXYD5 is also highly expressed in the salivary gland, tonsil and stomach, whereas they are absent in the same mouse tissue.
Similarly, murine FXYD5 exhibited the highest expression in the pineal gland, which produces melatonin and has been reported to exhibit oncostatic properties in tumor models of epithelial origin [138, 139]. Given the correlation of increased
FXYD5 expression and tumor metastatic potential, an inverse relationship between FXYD5 expression and melatonin production in carcinoma development may be hypothesized.
This hypothesis may be supported by the observation that other FXYD family members, such as FXYD3, express different isoforms based on the differentiation state of a particular cell [140]. Two isoforms of FXYD3 were expressed during differentiation of CaCo-2 cells, with a long version expressed in the undifferentiated state and a short form expressed in differentiated, polarized cells. Unigene profiles show a low level of FXYD5 expression in bone marrow and the thymus, but a much stronger signal was observed in blood and in lymph nodes. It has been postulated the FXYD5 has a role in cell-cell adhesion, and our immunohistochemistry and immunoflourescence results support the notion that
FXYD5, much like the β-subunit of the Na,K-ATPase, may have a secondary role
37 as an adhesion molecule. FXYD5, which has a large, extracellular, mucin-like domain, may therefore act as an intercellular adhesion molecule for mature lymphocytes or terminally differentiated squamous epithelia. Tissue and species specific differences may also be the result of different transcription elements present in the FXYD5 promoter, although this has not yet been determined.
Studies using traditional, non-array methods, such as Northern blotting and quantitative RT-PCR, have found similar expression patterns of mouse and human FXYD5. Our northern blot and quantitative RT-PCR data support the notion that Fxyd5 is strongly expressed in the lung, an observation reinforced by quantitative RT-PCR. Although immunoblot analyses of murine tissue indicate that FXYD5 is more ubiquitously expressed, it is clear that FXYD5 is expressed in the lung. Much like the differences observed when comparing Unigene expression profiles, tissue-specific post-translational modifications may affect the stability of the mature protein and account for transcription versus translation patterns between tissues. For example, FXYD3 from both species associates with and modulates the Na,K-ATPase, mouse FXYD3 has two transmembrane domains and lacks a cleavage signal whereas human FXYD3 has only 1 transmembrane domain and is a type I transmembrane protein [140].
Alternatively spliced transcripts have also been observed in the testes and brain, and a recent study has identified two human C-terminus FXYD5 isoforms, suggesting an additional level of complexity in the modulation of tissue-specific
Na,K-ATPase activity [120]. This may significantly affect regulation of FXYD5
38 expression, similar to differences observed between human and chimpanzee conserved genes [141, 142].
Human and mouse FXYD5 may be fundamentally different proteins.
Although the human and mouse genomes have a similar number of protein coding genes (approximately 30,000), these genomes diverged approximately
65-110 million years ago [143]. A comparison of over 12,000 orthologues identified FXYD5 as one of the top 50 mouse genes to have undergone rapid evolutionary changes, with high a KA/KS ratio of 0.66 (see footnote 1) [144]. A possible result of gene duplication and fixation, FXYD3, 1, 7 and 5 are genomic neighbors on mouse and human chromosomes 7 and 19, respectively. Such differences are observed in their migration patterns on SDS-PAGE. While each protein has a similar number of glycosylation sites and the predicted molecular weight is almost the same, the large difference in isoelectric point and charge at pH 7.0 apparently shifts their electrophoretic mobility, resulting in mature human and mouse FXYD5 proteins of 37 and 24 kDa, respectively. The migration patterns of chimeric proteins suggest that the C-terminus, which in human
FXYD5 has a large number of negative charges, is the critical determinant governing this difference, which is unique among FXYD family members. Using similar methods, others have reported that the extracellular domain mediates the effect of FXYD proteins on the apparent ATP affinity, and structural studies have demonstrated that the transmembrane domain regulates the effects on the Na,K-
ATPase cation affinities [88, 89, 145-147]. This may be particularly relevant for
39 understanding how Na,K-ATPase activity is modulated by FXYD5, which has a very short intracellular domain and is the most divergent FXYD family member.
Footnotes:
1 KA/KS is defined as ratio of is the ratio of non-synonymous (amino-acid changing) to synonymous (silent) substitution rates in protein-coding genes. To calculate the KA/KS ratio, the codons of two protein coding DNA sequences are aligned according to the amino acid pairwise alignment. KA is then estimated from the numbers of non-synonymous (amino acid replacing) substitutions at each non-synonymous site. Similarly, KS is estimated from the numbers of synonymous substitutions per synonymous site. If - KA ≅ KS, little or no selection has occurred among different sites; - KA << KS, strong purifying selection has reduced the fixation rate of deleterious mutations; and - KA > KS, the amino acid change fixation rate is higher than the neutral substitution rate. This is direct evidence for positive selection of amino acid substitutions which offer a selective advantage to the organism [144].
40
FXYD5 Human Mouse Genbank acc. # NM_014164 NM_008761 Chromosome 19q12-q13.1 7B1
Length (a. a.) 178 178 Size (kDa) 19.47 19.45 % Homology (a. a.) 100 54.5 Charge at pH 7.0 -3.9 4.01 Isoelectric point 5.51 9.13 Glycosylation sites 28 24 mRNA length (bp) 890 1144 # of ESTs 390 193 total sequences1 6808401 4204718 total clusters 123808 79710
Table 2-1. Comparison of human and mouse FXYD5. Human and mouse sequences were obtained from Genbank and analyzed using Vector Nti
(Invitrogen, Carlsbad, CA). 1 total sequences current as of 29-Oct-07 for mouse and 17-Nov-07 for human.
41 Figure 2-1. Unigene profile of human FXYD5 tissue expression. Sequence data from Homo sapiens, Unigene Hs. 333418, containing 123,808 clusters of
6,908,401 total sequences through 17-November-2007. Transcript sequences originated from the same transcription locus, demonstrating tissue specific expression patterns. TPM, transcripts per million; intens, spot intensity based on
TPM; Gene, Gene expressed sequence tag (EST); Total EST, total expressed sequence tags in pool.
42 Figure 2-1
Tissue TPM Intens Gene/Total EST adrenal gland 60 2 / 33320 ascites 124 5 / 40067 bladder 99 3 / 30133 blood 120 15 / 124121 bone 41 3 / 71794 bone marrow 101 5 / 49155 brain 16 18 / 1104170 cervix 0 0 / 48501 connective tissue 86 13 / 149627 ear 0 0 / 16343 embryonic tissue 41 9 / 215806 esophagus 0 0 / 20292 eye 47 10 / 210747 heart 44 4 / 90308 intestine 89 21 / 235308 kidney 18 4 / 212568 larynx 0 0 / 24435 liver 19 4 / 208302 lung 79 27 / 338047 lymph 22 1 / 44399 lymph node 163 15 / 91858 mammary gland 90 14 / 154293 muscle 9 1 / 108148 nerve 0 0 / 15765 ovary 19 2 / 102646 pancreas 116 25 / 215273 parathyroid 0 0 / 20633 pharynx 96 4 / 41490 pituitary gland 0 0 / 16734
43 Figure 2-1 continued placenta 38 11 / 283906 prostate 36 7 / 190868 salivary gland 197 4 / 20295 skin 132 28 / 210888 soft tissue 76 1 / 13146 spleen 55 3 / 54059 stomach 164 16 / 97138 testis 6 2 / 331289 thymus 0 0 / 81256 thyroid 20 1 / 47933 tongue 15 1 / 65977 tonsil 176 3 / 17031 trachea 19 1 / 52430 umbilical cord 145 2 / 13765 uterus 47 11 / 233915 vascular 77 4 / 51930
44 Figure 2-2. Unigene profile of murine Fxyd5 tissue expression. Sequence data from Mus musculus, Unigene Mm. 1870, containing 79,710 clusters of
4,204,718 total sequences through 28-October-2007. Transcript sequences originated from the same transcription locus, demonstrating tissue specific expression patterns. TPM, transcripts per million; intens, spot intensity based on
TPM; Gene, Gene expressed sequence tag (EST); Total EST, total expressed sequence tags in pool.
45 Figure 2-2.
Tissue TPM Intens Gene/Total EST adipose tissue 0 0 / 1576 adrenal gland 0 0 / 2343 bladder 0 0 / 15093 blood 580 10 / 17217 bone 59 2 / 33629 bone marrow 21 3 / 139698 brain 12 6 / 474042 connective tissue 50 1 / 19956 dorsal root ganglion 0 0 / 11284 embryonic tissue 19 13 / 678403 epididymis 0 0 / 2991 extraembryonic tissue 94 7 / 74220 eye 5 1 / 187857 fertilized ovum 0 0 / 27442 heart 0 0 / 53749 inner ear 26 1 / 38361 intestine 34 3 / 85851 joint 116 2 / 17205 kidney 0 0 / 124074 liver 0 0 / 110468 lung 30 3 / 99576 lymph node 131 2 / 15224 mammary gland 26 8 / 306321 molar 0 0 / 3630 muscle 0 0 / 27490 nasopharynx 0 0 / 8103 olfactory mucosa 0 0 / 3353 ovary 0 0 / 54820 oviduct 0 0 / 3640
46 Figure 2-2 continued pancreas 0 0 / 106585 pineal gland 764 3 / 3926 pituitary gland 0 0 / 17544 prostate 0 0 / 30012 salivary gland 0 0 / 19477 skin 244 29 / 118673 spinal cord 0 0 / 23855 spleen 31 3 / 95057 stomach 0 0 / 30554 sympathetic ganglion 0 0 / 9637 testis 0 0 / 118110 thymus 74 9 / 121546 thyroid 112 1 / 8891 tongue 194 2 / 10284 turbinate 0 0 / 1389 uterus 288 2 / 6933 vagina 0 0 / 5946 vesicular gland 0 0 / 2192
47 Figure 2-3
A
B
FXYD5
28S
Figure 2-3. Tissue distribution of murine Fxyd5 mRNA expression. (A)
Densitometry of murine Fxyd5/28S tissue expression shown in (B); Upper panel:
Northern blot of 10 µg of total RNA prepared from mouse tissue, probed with a
365 bp fragment of murine FXYD5 cDNA or Lower panel: 28S RNA stained with
methylene blue.
48
Figure 2-4
Figure 2-4. Tissue specificity of murine FXYD5. Quantitative RT-PCR analysis demonstrating that Fxyd5, normalized to Gapdh, is increased 5-fold in total lung tissue preparations from C57BL/6J mice.
49 Figure 2-5
1 2 3 4 5 6 7 M 8 9 M2
2.0 kb
1.5
0.8 1.0 Kb 0.8 0.65 0.5 0.4 0.4 0.2 0.3 0.1 0.2
Figure 2-5. FXYD5 is ubiquitously expressed in multiple cell lines. RT-PCR of FXYD5 from human, mouse and canine cell lines demonstrating prominent
FXYD5 expression. Lanes 1, HEK 293; 2, Calu-3; 3, NIH 3T3; 4, A549; 5, MDCK;
6, MEF; 7, HuH7; 8, ΔF508 mouse lung; 9, PLC/PRF5; M and M2, DNA markers.
Arrow, 556 bp FXYD5 RT-PCR product.
50 Figure 2-6
A B
M 1 M 1 2 200 200 116 116 97 97 66 66
45 45
31 31
21 21
14 * 14 10 10 *
Figure 2-6. FXYD5 antibody 562 recognizes murine FXYD5. (A)
Representative immunoblot of 10 µg crude membrane preparations from murine airway LA4 cells demonstrating FXYD5 562 antisera recognizes mature FXYD5
(arrow, Lane 1). (B) Immunoblot of 50 µg membrane preparations from LA4 cells transfected with pKCERmF5kFlag and probed with anti-M2 Flag antibody demonstrating that FXYD5-Flag is approximately 27 kDa (arrow, Lane 2) compared to control transfected LA4 cells (Lane 1). *, immature murine FXYD5.
51
Figure 2-7. (A) Image of 6 week old C57BL/6J mouse skin tissue stained with anti-FXYD5 562 antisera or (B) non-specific control antibody. The sections were developed with 3,3’-diaminobenzidine (DAB) using DAB substrate kit (Vector
Laboratories) and counterstained with hematoxylin (Themo Electron Corporation,
Waltham, MA). Arrows (white) highlight FXYD5 stained cell membrane.
52
Figure 2-7 A
B
53 Figure 2-8
M 1 2 3 4 5 6 7 8 9
31
21
Figure 2-8. Observed mobility of murine FXYD5 is 27 kDa. Representative
immunoblot demonstrating endogenous FXYD5 tissue expression in 50 µg crude
membrane preparations from C57BL/6J mice using antibody 562. Lane 1, liver;
2, lung; 3, heart; 4, brain; 5, kidney; 6, small intestine; 7, skeletal muscle; 8,
spleen; 9, pancreas; M, protein markers.
54 94 GP ET DP DP
S T VSDP V V 1 R R R 18 H C NR NR NR L R C C C
TKMATSNP EGTDGP-L QF RL 80 0 TQQL TQQL TQQL 17 E Q junction Chimera QT QT QT
TAA --P Q 70 ). ). Q Q 0 16 REEATGS WPADETP T T T black bar black 60 PLIW SPTP 0 T T T 15 T P Q Q LQ V L TLRKRGLLVAAVLFITGIIILTSGKCRQLS TLRKRGLLVAAVLFITGIIILTSGKCRQLS TLRKRGLLVAAVLFITGIIILTSGKCRQLS 50 T H QTSPG AVYTE D E 0 PD PD PD 14 YD YD Y F
Human Human and mouse cDNA sequences were 9 PFFYDD - PF PF
NN DD E E E 40 ighlighted in blue, methionine residues highlighted in 0 SATTRDNVPD IMDIQVPTRA 13 LDSN SGFH Figure2 T T T P P P Q S H K AD AD SAD T S 30 IF SS TS TS TS 0 12 KP DT K K K KNFMPPSYIE TDVQTDPQTL P L SPS SPS SPS 20 RGQT RGQT PL RP S T E E E 0 11 LILP LILP A G AVSRI--- TDDTTTLS 10 P P P T H 0 G A A 10 K A RLCLLTIVALILPSRGQT RLCLLTIV RLCLLTIV K K S G S T S S S S S SSK L P T K 95 H H H 1 MS MS MS ) ) ) ) 9. Alignment of human and mouse FXYD5. 5) 5) 2) 5) - (1 (1 (1 (1 (9 (9 (9 (9
Figure 2 obtained from Genbank residues highlighted (accession in yellow, conserved residues numbers,h NM_014164, red. chimericfor and Splice usedjunction is indicated ( humanmouse constructs human; NM_008761, mouse). Identical D5 D5 us D5 D5 us XY XY XY XY Consensus Consensus ens ens e F n F ns e F n F ns MouseFXYD5 MouseFXYD5 HumanFXYD5 HumanFXYD5 us ma Co us ma Co mo hu mo hu
Figure 2-10. Schematic diagram of human and mouse FXYD5 Flag-tagged and chimeric constructs. FXYD family proteins share a conserved 35-residue domain that contains the FXYD domain (bold), a transmembrane region
(overlined) that includes two conserved glycines and a conserved cytoplasmic serine residue (5,12). (A) Schematics for chimeric constructs are as follows: human (solid bar), mouse (cross-hatched), Flag-tag (horizontal bars). (B) Lanes
1-6, 10 µg transiently transfected HEK 293 cells; 7-12, 50 µg transiently transfected LA4 cells. Lanes 1 and 7 are 293 and LA4 vector transfected controls, respectively. Arrows denote bands corresponding to use of internal in- frame ATG sites. (C, D) HEK293 cells transiently transfected with pKCERhF5kQ22Flag. After 24 hr, cells were fixed, stained for nuclei (blue) and the Flag epitope (red) and visualized on a Zeiss 200M Axiovert inverted microscope using the 63X objective. Scale bar = 20 µm.
56 Figure 2-10
A PFXYD G G S Lane
pKCERhF5kFlag 2
pKCERhF5kQ22Flag 3
pKCERhF5kQ22FlagmF5 4,10
pKCERmF5khF5Flag 5,11
pKCERhF5kmF5Flag 6,12
pKCERmF5kFlagtrunc 8
pKCERmF5kFlag 9
B
C D
57
Chapter 3: FXYD5 modulates Na+ absorption and is increased in cystic fibrosis airway epithelia
Timothy J. Miller1,2 and Pamela B. Davis2 From the Departments of 1Pharmacology and 2Pediatrics, School of Medicine, Case Western Reserve University, Cleveland, Ohio
Abstract
FXYD5, also known as Dysadherin, belongs to a family of tissue-specific regulators of the Na,K-ATPase. We determined the kinetics effects of FXYD5 on
Na,K-ATPase pump activity in stably transfected MDCK cells. FXYD5 significantly increased the apparent affinity for Na+ 2-fold, and decreased the
+ apparent affinity for K by 60% with a 2-fold increase in Vmax(K+), a pattern that would increase activity and Na+ removal from the cell. To test the effect of increased sodium uptake on FXYD5 expression, we analyzed MDCK cells stably transfected with an inducible vector expressing all three subunits of the epithelial sodium channel (ENaC). Na,K-ATPase activity increased 6-fold after 48-hour
ENaC induction, but FXYD5 expression decreased 75%. FXYD5 expression was also decreased in lung epithelia from mice that overexpress ENaC, suggesting that chronic Na+ absorption by itself downregulates epithelial FXYD5 expression.
Patients with cystic fibrosis (CF) display ENaC-mediated hyperabsorption of Na+ in the airways, accompanied by increased Na,K-ATPase activity. However,
FXYD5 was significantly increased in the lungs and nasal epithelium of CF mice as assessed by RT-PCR, immunohistochemistry and immunoblot analysis
(P<0.001). FXYD5 was also upregulated in nasal scrapings from human CF
58 patients compared to controls (P<0.02). Treatment of human tracheal epithelial
(HTE) cells with a CFTR inhibitor (CTFRinh-172) confirmed that loss of CFTR function correlated with increased FXYD5 expression (P<0.001), which was abrogated by an inhibitor of NF-κB. Similarly, stimulation of NF-κB activity with the pro-inflammatory cytokines TNFα/IL-1β upregulated FXYD5 expression and was blocked by a separate chemical inhibitor of NF-κB. Thus FXYD5 is upregulated in CF epithelia and this change may exacerbate the Na+ hyperabsorption and surface liquid dehydration observed in CF airway epithelia.
59 Introduction
The FXYD protein family, typified by the γ-subunit (FXYD2) of the Na,K-
ATPase, is identified by a signature 35-residue domain containing an invariant, extracellular PFXYD sequence [45, 87]. The mammalian family contains 7 members, many of which have been shown to be tissue specific subunits of the
Na,K-ATPase responsible for fine tuning its kinetic behavior in response to extracellular signals (reviewed in [86, 96]). While the primary function of the
Na,K-ATPase is to maintain intracellular sodium/potassium homeostasis, decreased expression of the α and β subunits has been correlated with neoplastic transformation [68, 70] and recent literature has implicated FXYD proteins in cancer progression [70, 105-107, 111-114, 116].
Originally identified as a gene induced in NIH-3T3 cells transformed with the oncoprotein E2a-Pbx1, FXYD5 was subsequently cloned and characterized as a cancer-associated cell membrane glycoprotein [103, 105]. FXYD5 encodes a 178 amino acid protein that includes a putative signal sequence, a single transmembrane domain and a short cytoplasmic tail. FXYD5 is unique in the
FXYD family, possessing a heavily O-glycosylated, extended extracellular domain [105, 135]. Transfection of FXYD5, also known as dysadherin, into liver cells led to decreased cell-cell adhesion correlated with diminished E-cadherin levels [105]. Elevated FXYD5 expression in tumors from patients with thyroid, esophageal, colorectal, stomach, cervical, pancreatic, testicular, head/neck or lung cancer correlates with a poor prognosis, suggesting that FXYD5 may be critical determinant regulating the role of the Na,K-ATPase in determining cell
adherence and polarity [106, 108, 110-115, 136, 148]. Indeed, knockdown of
FXYD5 expression correlates with decreased cell motility [107, 135], independent of E-cadherin expression, and recent evidence indicates that murine FXYD5 interacts with and may regulate the Na,K-ATPase in Xenopus oocytes [119, 120].
However, it is unclear how FXYD5 affects the functional properties of Na,K-
ATPase pump activity.
FXYD5 is highly expressed in the basal layer of squamous epithelia, endothelia and lymphocytes, as well as in ion transport tissues such as the kidney and lung [105, 113, 135, 136], where modulation of the Na,K-ATPase may be critical for specialized functions such as sodium reabsorption. In lung epithelial cells, the Na,K-ATPase is critical for maintaining transepithelial sodium transport and maintaining the alveolar fluid absorption necessary for efficient gas exchange. Thus, we investigated whether FXYD5 expression is altered in epithelial cell and animal models that exhibit altered transepithelial sodium transport.
Cystic fibrosis (CF) is a common genetic disease among Caucasians, caused by lesions in they cystic fibrosis transmembrane conductance regulator gene, CFTR, resulting in the inability to transport chloride through the apical membrane, particularly in epithelia lining the airway. A hallmark of CF is the hyperabsorption of sodium through the epithelial sodium channel (ENaC) [36]. In the airway, increased sodium uptake is postulated to dehydrate the pericilliary layer, impair mucociliary clearance, cause mucus buildup in the lung, and lead to bacterial trapping. The increased bacterial burden causes infection, inflammation
61 and tissue damage characteristic of CF lungs [36]. Since it has been shown that airway epithelia from CF patients have increased Na,K-ATPase pump number and activity compared to normal subjects [32, 42, 149], we investigated whether
FXYD5 is altered in the airway epithelia of CF mice and human systems and examined how FXYD5 may alter transepithelial sodium transport along the
ENaC/Na,K-ATPase axis.
Materials and Methods
Cell lines. The mouse lung epithelial cell line LA4, human embryonic kidney
(HEK) 293 and Madin-Darby canine kidney (MDCK) cells were obtained from the
American Type Culture Collection (Manassas, VA). LA4 cells were grown in
Kaighn’s modification of F12 medium (F12K, Mediatech, Inc., Herndon, VA), HEK
293 cells were grown Earle’s modification of MEM media (Mediatech) and MDCK cells were grown in 1:1 (vol:vol) DMEM/F-12 media (Mediatech). An MDCK cell line stably transfected with an inducible vector expressing the α, β and γ subunits of the epithelial sodium channel (ENaC), a generous gift of Dr. Calvin Cotton
(Case Western Reserve University, Cleveland, OH), has been previously described (MDCK clone 29.1 [150, 151]) and is referred to below as the MDCK-
ENaC cell line. ENaC expression was induced by the addition of 1 µM dexamethasone and 2 mM sodium butyrate to serum-free MDCK culture media and analyzed 36-48 hours later. All media were supplemented with 10% heat- inactivated FBS and all cells were grown in a 37°C, 5% CO2-95% 02 atmosphere.
62 Mouse strains. Breeding pairs of heterozygote congenic mice (>N10) bearing the
S489X mutation (B6.129P2-Cftrtm1unc, stock no. 2196) and C57BL/6J mice were purchased from Jackson Laboratory (Bar Harbor, ME). Breeding pairs of heterozygote mice bearing the ΔF508 Cftr mutation in mixed genetic background were a kind gift from Dr. Kirk Thomas from the University of Utah, and were backcrossed into the C57BL/6J background for at least 10 generations in the CF
Animal Core Facility at CWRU before use. Cystic fibrosis mice for these strains are indicated by their Cftr mutation and are referred to as “CF mice”. Breeding pairs of heterozygote mice bearing the sodium channel, nonvoltage-gated 1 beta,
Scnn1b, under control of the rat secretoglobin, family 1A, member 1, Scgb1a1 promoter (B6;C3H-Tg(Scgb1a1-Scnn1b)6608Bouc/J, stock number 005315) were purchased from Jackson Labs (Bar Harbor, ME). These mice are on the
B6C3F1/J background and are a F1 hybrid cross of C57BL/6J and C3H/HeJ mice. All animal studies were performed under CWRU IACUC approved protocols.
Human nasal scrapes. CF patient and control nasal epithelium were obtained as previously described [152]. 3-4 scrapes from one female and 3 male CF patients, age 18-42 years, and 6 female, 1 male normal volunteers, age 26-53 years were obtained and immediately stored at 4ºC in RNAlater (Qiagen, Inc., Valencia, CA) for processing. Parallel scrapes were obtained and scored for percentage epithelial cells present, which was used to normalize sample data. The protocol was approved by the Institutional Review Board (IRB).
63
HTE cells and treatment with CFTRinh-172. Human tracheal epithelial (HTE) cells were recovered from necropsy specimens as previously described [153] under an exempt IRB protocol. Multiple donors were used for analysis. Briefly, HTE cells were grown in an air-liquid interface (ALI) on collagen-coated, semi permeable membranes (1x106 cells / 1 cm2 filter, transwell-clear polyester membrane, Costar, Corning, N.Y.) as previously described [153, 154] and allowed to differentiate in serum-containing media for three or four weeks. Then, on day 0, cells were switched to submerged culture in serum-free media and treated with either DMSO 1:1000 (vehicle control, normal cells, Sigma, St. Louis,
MO), or 20 µM CFTRinh-172 (a kind gift of Alan Verkman, UCSF), prepared in
DMSO, and diluted from a 1:1000 stock. Similarly, the NF-kB inhibitor pyrrolidinecarbodithioate (PDTC, Sigma, St. Louis, MO) was diluted in serum- free media to a final concentration of 0.1 mM and was added simultaneously with
CFTRinh-172. Drugs were added to both the basolateral and apical media, which were replenished every 24 hours. Cells were isolated for analysis following 3 days of drug treatment.
HTE were stimulated for 1 hour at 37ºC with 10 ng/ml tumor necrosis factor-alpha (TNF-α) and 5 ng/ml interleukin 1-beta (IL-1β) (Sigma, St. Louis,
MO) in 200 µl Hanks balanced salt solution (HBSS, Stratagene, Inc., Cedar
Creek, Texas) added to the apical surface. NF-κB activation was inhibited by pretreating HTE with 20 µM Bay inhibitor 11-7085 (a kind gift of John Mieyal,
Case Western Reserve University, Cleveland, OH) for 1 hour. After stimulation,
64 the apical surface was washed twice with HBSS, the air-liquid interface re- established and the cells isolated for RNA preparation 4 hours later.
RNA isolation. Total RNA was isolated from 6 to 8 week old mice, 10 cm dishes of LA4, HEK293, MDCK cells or HTE cultures using the RNAprotect kit according to the manufacturer’s instructions (Qiagen, Inc., Valencia, CA) and stored at –
80ºC. RNA was quantified on a spectrophotometer and visualized by agarose gel electrophoresis to determine quality.
RT-PCR, cloning and site-directed mutagenesis of FXYD5. The Superscript II
One-Step RT-PCR kit (Invitrogen, Carlsbad, CA) was used to reverse transcribe and amplify FXYD5 from HEK293 cells. FXYD5 cDNA was isolated by RT-PCR using primers designed from accession numbers NM014164 (human) and
NM008761 (mouse), which contained a HindIII and NotI restriction site on the 5’ and 3’ end respectively. The following primers were used to generate FXYD5 clones: Human 5’AAGCTTGCTAGCGCCGCCACCATGTCGCCCTCTGGTC
GCCTGTGTCT (forward), 5’ AGTCGTCTAGATCACCTGCAACGATTC
CGGCATAAC (reverse); mouse 5’ AAGCTTGCTAGCGCCGCCACCAT
GTCACTGTCCAGTCGCCTGTGTCT (forward) 5’ AGTCGTCTAGATC
ACCTGTGGCGATTCAGGCAAATT (reverse). RT-PCR was performed as follows: reactions were incubated at 50ºC for 30 minutes, followed by 2 minute initial denaturation at 95ºC and 40 cycles of 94ºC, 1 minute denaturation, 1 minute 58ºC primer annealing, and 45 seconds of primer extension. RT-PCR
65 products were digested with HindIII and NotI restriction enzymes and agarose gel purified using the Qiaquick gel purification kit (Qiagen, Inc., Valencia, CA). cDNAs were subcloned in pBSK2 vector to create pBhF5k (human) and pBmF5k
(murine). To generate an N-terminus Flag tag in human FXYD5 that did not alter the N-terminus signal sequence, codon Q22 was mutated using the Quickchange site-directed mutagenesis kit (Stratagene, Inc., Cedar Creek, Texas). A silent mutation was introduced (CAG mutated to CAA) to create a new restriction site,
Acl1. The following sequence was then inserted in frame at the Acl1 site to produce pBhF5kQ22Flag: 5’ CGGATTACAAAGATGATGATGATAAGA 3’. The original and modified versions of FXYD5 were then subcloned into the pTracerCMV2 plasmid (Stratagene, Inc., Cedar Creek, Texas, Inc.) using the
EcoRV/Not restriction sites to create the pThF5kQ22Flag vector used for stable cell line expression. All cDNA sequences and mutations were verified by double- strand DNA sequencing.
Creation of MDCK-hF5Flag stable cell line. MDCK cells were transfected in 10 cm dishes with vector (sham) or pThF5kQ22Flag using Fugene 6 (Roche
Molecular Biochemicals, Mannheim, Germany) according to the manufacturer’s instructions to create the MDCK-sham and MDCK-hF5Flag cell lines. Each line was selected and maintained in 400 µg/ml zeocin and analyzed after 3 weeks incubation by fluorescence activated cell sorting (FACS) using the BD-FACSAria
(BD Biosciences, San Jose, CA) at 488 nM to obtain multiple positive clones.
Positive clones were identified by immunoblot and immunofluorescence analysis
66 using the antibodies directed against the Flag epitope (Sigma, St. Louis, MO and
Bethyl Labs, Inc, Montgomery, TX) and the α1-Na,K-ATPase (clone 464.6,
Millipore, Inc., Billerica, MA).
Quantitative RT-PCR. 0.5 µg of total RNA was utilized to synthesize cDNA with the First Strand cDNA synthesis kit (Roche Molecular Biochemicals, Mannheim,
Germany). cDNA synthesis was performed using oligo-p(dT)15 primer per kit instructions and stored at -20ºC. Quantitative PCR was performed using the
Lightcycler FastStart DNA master Sybr green kit (Roche Molecular Biochemicals,
Mannheim, Germany). The mouse and canine sequences are conserved across the primer sequences used in this study. The primer sequences used for human and mouse FXYD5 are: human 5’ ATGTCGCCCTCTGGTCGCCTG (forward), 5’
TCACCTGCAACGATTCCGGCA (reverse) and mouse 5’ ATGTCACTGTCCAGT
CGCCTGTGTCTCCTCACT (forward), 5’ TCACCTGTGGCGATTC
AGGCAAAATTGAGACAA (reverse). DNA was amplified using a Roche
Lightcycler with the following parameters: 95ºC 10 minute initial denaturation followed by 35 cycles of 95ºC 10 second denaturation, 58ºC 7 second anneal,
72ºC 10 second extension. Copy number was measured against a standard curve of linearized plasmid DNA and normalized to GAPDH. Second point derivatives were used for PCR analysis. Specificity of DNA amplification was assessed by melting point analysis and confirmed by agarose gel electrophoresis.
67 Immunohistochemistry of mouse tissue. Adult ΔF508 mice and their wild-type littermates were sacrificed and the lungs fixed by perfusion using 2% paraformaldehyde in 1x PBS (PH 7.4) via the left ventricle. Lung tissue was immersed in the same fixative solution for 48 hrs at room temperature, dehydrated and embedded in paraffin using standard histological procedures. 5
µm sections were mounted onto Fisherbrand superfrost slides coated with
Vectabond (Fisher 12-550-15). After deparaffinization, the sections were treated with cold methanol for 15 minutes followed by 1% hydrogen peroxide in distilled water for 30 minutes at room temperature to block endogenous peroxidase activity, washed with PBS and incubated with 1.5% normal goat serum/PBS
(Vector Laboratories, Burlingame, CA, PK-6101) for 4 hrs at room temperature.
The sections were then incubated with 6.8 µg/ml of affinity purified 562 polyclonal rabbit antibody against FXYD5 overnight at 4ºC. Blocking solution without primary antibody was used as a negative control. After washing 3 times in 1x
PBS, the sections were incubated with 1.5% biotinylated goat anti-rabbit IgG antibody 1 hr at room temperature. After washing 3 times with PBS, the sections were incubated with Vectastain ABC reagent for 30 minutes and washed again.
The sections were developed with 3,3'-diaminobenzidine (DAB) using DAB substrate kit (Vector Laboratories) for 5 minutes. After washing 2 times with water, the sections were counterstained with hematoxylin (Themo Electron
Corporation, Waltham, MA).
68 Isolation of crude membranes. Crude membranes were prepared as previously described [119]. Mouse tissue or cultured cells were isolated, washed with PBS, suspended in 25 mM imidazole, 1 mM EDTA, 250 mM sucrose and protease inhibitor cocktail (Sigma, St. Louis, MO) and manually homogenized with 35 strokes of a prechilled Dounce homogenizer. Nuclei, unbroken cells and mitochondria were separated by centrifuging at 6000 x g for 5 minutes. The supernatant was collected and saved, while the pellet was homogenized again and centrifuged. The two supernatants were combined and centrifuged at
125,000 x g to pellet crude microsomal membranes. The pellets were resuspended in 25 mM imidazole, 1 mM EDTA, 10 mM RbCl and stored at 4ºC.
Indirect immunofluorescence. Cells were cultured in 4-well chamber slides
(Permanox), washed twice with phosphate-buffered saline (PBS) and fixed for 5 minutes in 100% ice-cold methanol. Slides were then rinsed twice with PBS, blocked for non-specific antibody binding by incubating the cells in 4% bovine serum albumin for 1 hour (Invitrogen, Carlsbad, CA). Primary rabbit anti-ECS
(Flag) antibody (2 µg/ml) (Bethyl Labs, Montgomery, TX) or mouse anti-α1 Na,K-
ATPase subunit antibody (clone 464.6, Millipore, Inc., Billerica, MA) (1 µg/ml) were diluted in 1% BSA/PBS, incubated on monolayers for 45 minutes and aspirated. Monolayers were washed twice with PBS and then incubated in 1%
BSA/PBS containing goat anti-mouse Alexa-Fluor 488 or goat anti-rabbit IgG
Alexa-Fluor 568 (1:250, Invitrogen, Carlsbad, CA) for 45 minutes. Cells were then washed twice, the nuclei counterstained with Hoechst 33342 dye
69 (Invitrogen, Carlsbad, CA) and mounted with Fluormount-G. Slides were allowed to dry overnight and immunofluorescent localization was assessed on a Zeiss
200M Axiovert inverted microscope, with a DG4 switchable fluorescent light source (Sutter Instrument Company, Novato, CA) and a 12-bit CoolSnap HQ camera (Roper Scientific, Tucson, AZ) under control of MetaMorph v 6.2
(Molecular Devices, Sunnyvale, CA. Images were obtained with a 63X numerical aperture 1.3 fluar lens using excitation and emission filter passbands of 260 ±
20nm and 645 ±30 nm, respectively. Typical exposure times for individual frames were 150 ms.
86Rb+ uptake transport assay. Unidirectional Rb+ influxes into cells were measured and calculated as previously described, using 86Rb+ as a congener of
K+ uptake, with minor modifications [48]. Briefly, were cells grown to confluence in 24-well tissue culture plates (Costar, Corning, NY) for 2-3 days. All solutions were maintained at 37°C. For Rb+ (K+) dependent 86Rb+ uptake, cells were washed twice in the following assay buffer (in mM): 140 choline chloride, 10
NaCl, 10 HEPES pH 7.4, 5 glucose, 1 MgCl2, 1 CaCl2, 3 BaCl2, 5 µM monensin and 50 µM bumetanide. For assays varying intracellular Na+ dependent 86Rb+ uptake, the wash and incubation solutions were similar except that 2 mM RbCl,
10 µM monensin and 2-70 mM NaCl were used with choline chloride added to maintain isotonicity. Cells were then preincubated in assay buffer in the presence or absence of 100 µM ouabain for 10 minutes at 37°C and washed twice. The assay was initiated by the addition of 25 µl (1-2 µCi 86RbCl) to 225 µl assay
70 buffer but with various concentrations of RbCl as indicated per well. The reaction was carried out for 6 minutes at 37°C during which the rate of 86Rb+ uptake remained constant (data not shown) [155] and the cells washed in assay buffer without monensin. Cells were allowed to air dry for an hour, solubilized in 500 µl
2% SDS, 0.2 mM NaOH overnight, and sampled to determine 86Rb+ uptake in a scintillation counter. Multiple hF5Flag positive clones were tested, with no significant difference in 86Rb+ uptake observed between clones. Ouabain- sensitive 86Rb+ uptake was calculated as the difference between total and ouabain-insensitive 86Rb+ accumulation and samples were normalized to protein content using the DC-Protein assay (Bio-rad).
Kinetic analysis. The data for Na+ and K+ dependent 86Rb+ (K+) influxes were analyzed by nonlinear regression using Prism 4 (Graphpad Software). All data were analyzed using a Michaelis-Menten model for either a non-cooperative two- site or three-site model that assumes identical noninteracting ligand binding sites as previously described [47, 48, 130, 156]:
n ν = Vmax/(1+Ks/[S]) (Equation 1)
+ where n is the number of sites and Ks is the apparent affinity for extracellular K
+ (Kk(ext)) or for intracellular Na (KNa(in)). Data were also analyzed according to a highly cooperative model:
ν = Vmax[S] n /K’ + [S]n) (Equation 2)
71 1/n + where K0.5 = K’ , and n is the number of sites for extracellular K (n=2) or cytoplasmic Na+ (n=3). Experimental data points were fit to Equations 1 and 2 to obtain the values of Vmax and apparent K0.5. Specificity constants were obtained using the relationship:
Specificity constant = Vmax(S)/K0.5(S) (Equation 3) where (S) represents the values obtained for Na+ or K+. Results from both
Equations 1 and 2 are presented in Table I, whereas the figures show curves that have been fit using Equation 1.
Statistics. Microsoft Excel was used for calculating Student’s T-test. Sigma Stat
(Systat Software Inc., San Jose, CA) and Prism 4 (Graphpad Software) were used to calculate ANOVA statistics using the Student-Neuman-Kuels regression analysis for pairwise comparisons. The Wilcoxen matched pairs test was used to calculate significance values for 1-substrate kinetic data obtained from 86Rb+ uptake experiments. P<0.05 was used to declare statistical significance unless otherwise noted.
72 Results
FXYD5 modulates Na,K-ATPase ion transport
Flag-FXYD5 is stably expressed at the cell membrane. In order to develop an epithelial cell culture model system to assess the kinetic properties of FXYD5 on the Na,K-ATPase, MDCK cells were stably transfected with pThF5kFlag or empty
(sham) vector and selected in zeocin. After 3 weeks, positive clones were identified by FACS analysis (data not shown), subcultured, and assessed for immunofluorescence reactivity to anti-Flag antibodies. Cells were fixed in methanol and immunostained for FXYD5-Flag and the α1-subunit of the Na,K-
ATPase. The Flag antibody reacted strongly to MDCK-hF5Flag cells and recognized membrane-localized mature human FXYD5, but not the empty vector
(sham) control (Figure 3-1A,D). Immunofluorescent staining of the α1-subunit of the Na,K-ATPase demonstrated that membrane-localization of pump subunits was unchanged in pThF5kFlag transfected cells (Figure 3-1C, 3-1F), and that
FXYD5-Flag co-localized with the Na,K-ATPase in the cell membrane (Figure 3-
1B, 3-1E). Immunoblot analysis of crude membrane preparations of MDCK- hF5Flag cells demonstrated that mature human FXYD5-Flag protein migrated as an indistinct band of approximately 35 kDa, indicating heavy glycosylation
(Figure 3-1G), and that surface expression of the Na,K-ATPase was similar to vector transfected cells (Figure 3-1H). In separate experiments, FXYD5-Flag was coimmunoprecipitated with the α subunit of the Na,K-ATPase and this subunit was coimmunoprecipitated with antibody to FLAG or to FXYD5 (data not shown).
Thus we developed an appropriate epithelial cell model for testing the kinetic parameters of FXYD5 and its effects on Na,K-ATPase pump activity.
FXYD5 alters Na,K-ATPase pump kinetics. The proteins of the FXYD family are believed to be responsible for fine-tuning the kinetics of Na,K-ATPase pump activity in a tissue-specific manner. However, while previous studies have shown that FXYD5 associates with the Na,K-ATPase, it is unclear how FXYD5 modulates pump activity. To assess this question, Na+ and K+ dependent 86Rb+
(a congener of K+) uptake were compared in MDCK-hF5Flag and vector control cells. Data were analyzed by fitting experimental data points to non-linear regression models based on Michaelis-Menten kinetics as originally described by
Garay and Garrahan [156] and are summarized in Table 3-1. Cells were incubated with 50 µM bumetanide and in the presence or absence of 100 µM ouabain to determine Na,K-ATPase specific 86Rb+ uptake. Figure 3-2A shows that FXYD5 significantly increased the K0.5(K+) by approximately 60% (K0.5 0.07 ±
0.01 mM for vector, 0.12 ± 0.01 for FXYD5 transfected, P<0.03) and increased the K+ dependent maximal Na,K-pump uptake 2-fold compared to vector transfected control MDCK cells (P<0.01). In contrast, FXYD5 decreased the
K0.5(Na+) 2-fold (K0.5 9.24 ± 2.68 mM for vector, 4.53 ± 1.54 for FXYD5 transfected cells, P<0.05) without significantly affecting the Na+ dependent maximum pump rate (Figure 3-2B). The specificity constant, Vmax/K0.5, was calculated and used to compare Na,K-ATPase pump activity in the presence or absence of FXYD5
(Table I), and demonstrates that FXYD5 increases the overall pump efficiency for
74 both Na+ and K+. These results indicate that, similar to FXYD4 (CHIF), FXYD5 increases Na,K-pump transport activity approximately 4 fold at physiologically low intracellular Na+ concentrations [78] by increasing the apparent affinity for
Na+ and increasing the maximal rate of K+ transport. Taken together, these data suggest that FXYD5 may be involved in the regulation of active Na+ reabsorption in ion transport tissues such as the lung and kidney.
FXYD5 is decreased during chronic Na+ absorption
FXYD5 is downregulated after ENaC activation in MDCK-ENaC cells. To investigate whether increased Na,K-ATPase activity affects FXYD5 expression, we utilized a stable cell line that expresses high levels of the α, β and γ ENaC subunits upon induction in serum-free media [151]. Previous studies have shown that induction of ENaC expression generated a large increase in amiloride- sensitive Isc, but the effect on Na,K-ATPase activity was not shown [150]. As expected, induction of ENaC expression increased the amount of Na,K-ATPase immunostaining in the membranes of induced compared to control cells (Figure
3-3A-D). Similarly, a significant, 6-fold increase in ouabain- and bumetanide- sensitive 86Rb+ uptake was observed in MDCK-ENaC cells after ENaC induction
(n = 10, P<0.0001), indicating a large increase in Na,K-ATPase pump activity
(Figure 3E). Interestingly, quantitative RT-PCR (Figure 3-3F) revealed a 75% decrease in FXYD5 expression in MDCK-ENaC induced cells (n = 6, P<0.0001).
This suggests that overexpression of ENaC subunits, which results in increased
75 sodium extrusion via the Na,K-ATPase, may downregulate FXYD5 expression in epithelia.
FXYD5 is decreased in the lungs of Scnn1b transgenic mice. To confirm these findings in an in vivo model, mice overexpressing the Scnn1b (ENaC β- subunit) were examined. These mice exhibit accelerated Na+ absorption in lower airway epithelia and demonstrate some of the features of airway pathophysiology observed in CF [36]. Quantitative RT-PCR analysis of lung tissue from Scnn1b- positive mice revealed a 40% decrease in Fxyd5 expression compared to control littermates (Figure 3-4A) (n = 4, p<0.0001). Immunoblot analysis of crude membrane preparations confirmed that FXYD5 expression was decreased in
Scnn1b-positive mouse lung tissue (Figure 3-4B). Due to the multiple cell types present in total lung preparations, these results may underestimate the actual downregulation of FXYD5 specifically in lung epithelia. Based on our kinetic analysis of FXYD5 effects on Na,K-ATPase pump activity and our biochemical data indicating that FXYD5 is negatively regulated in vitro and in vivo by Na+ hyperabsorption, we speculated that FXYD5 expression would be decreased in
CF airway epithelia.
FXYD5 is upregulated in cystic fibrosis airway epithelia.
FXYD5 is upregulated in the nasal epithelia of CF mice. CF murine nasal epithelium is the airway tissue that most closely approximates the ion transport abnormalities observed in human CF lung and nasal epithelia, including ENaC
76 upregulation. In addition, the nasal epithelium of mice can be dissected from the underlying tissues and studied in isolation. Therefore, we utilized nasal epithelium from mice homozygous for the S489X mutation in cftr for analysis of
FXYD5 mRNA and protein expression. Quantitative RT-PCR analysis of nasal epithelia from S489X-/- mice revealed a significant (P<0.001), nearly three-fold increase in FXYD5 transcription compared to epithelia from wild-type littermates
(Figure 3-5A). To test whether this increase in FXYD5 mRNA transcription was accompanied by an increase in FXYD5 protein, we developed a polyclonal antibody that recognized the mature (highly glycosylated) form of murine FXYD5 at approximately 27 kDa (Supplemental data). Immunoblot analysis of membrane fractions from nasal epithelia confirmed an increase in mature FXYD5 expression from S489X-/- CF mice compared to wild type littermates (Figure 3-5B).
FXYD5 is increased in lungs of CF mice. Upper and lower airway epithelia from the lungs of ΔF508-/- CF mice were retrieved for immunohistochemical analysis. Similar to our observations in the nasal epithelia of S489X-/- CF mice,
FXYD5 expression was increased in the upper airway epithelia of ΔF508 CF mice compared to wild type littermates (Figure 3-6A,B). Interestingly, FXYD5 immunoreactivity was observed within the cytoplasm as well as at the cell surface in upper airway epithelia, similar to the pattern of tumor and non-tumor staining in squamous epithelia observed by us and others [105, 111]. Although
FXYD5 was increased in CF mice in the lower airway epithelium, here it appeared to localize to the cell membrane (Figure 3-6D, arrow) indicating a
77 possible cell-type specific mechanism governing membrane insertion. No- primary-antibody control tissue staining demonstrates no staining (Figure 3-
6E,F). Similar results were observed in lung tissue from S489X-/- mice (data not shown). Immunoblot analysis performed on crude membrane fractions from
ΔF508-/- CF lung tissue confirmed that FXYD5 is increased in the CF lung
(Figure 3-6G).
CFTR inhibition upregulates FXYD5 in human epithelia. To test whether increased FXYD5 expression also occurs in human CF epithelium, we obtained nasal scrapings from CF and control patients and performed quantitative RT-PCR for FXYD5 mRNA. FXYD5 is increased in nasal epithelia from CF patients (P<0.02, Figure 3-7A) compared to controls. We also studied human airway epithelial cells cultured at the air-liquid interface and nasal scrapes from control and CF patients. Cells from the same donor grown in this manner with and without the inhibitor are well matched for other genetic and environmental influences and constitute a valid test of the effect of lack of CFTR function in isolation from other factors. Previous studies have shown that treatment of non-CF human tracheal epithelial (HTE) cells grown on filters at the air-liquid interface in the presence of the CFTR inhibitor, CFTRinh-172, for 72 hours resulted in markedly reduced CFTR activity, no increase in ENaC activity, and increased inflammatory response [154]. HTE cells treated for three days with
CFTRinh-172 display a significant (P<0.001), 40% increase in FXYD5 expression compared to control cultures (Figure 3-7B), and this increase was completely
78 abrogated by treatment with the NF-κB inhibitor, pyrrolidinecarbodithioate
(PDTC, P<0.01) (Figure 3-7B). Similarly, stimulation of HTE cells with TNF-α/IL-
1β for 1 hour increased FXYD5 expression approximately 60% (P<0.001) after 4 hours, which was blocked by pretreatment with the NF-κB inhibitor BAY 11-7085
(P<0.01) (Figure 3-7C). Therefore, FXYD5 is increased in human CF airway epithelia as well as in CF mice, possibly due to alterations in pro-inflammatory signaling observed in the CF airway.
79 Discussion
Taken together with previous studies, our results indicate that FXYD5 specifically associates with the Na,K-ATPase and increases the catalytic efficiency of the pump for Na+ and K+. Similar to other reports, we observed that
FXYD5 increased Na,K-ATPase pump activity [120]. However, our 86Rb+ uptake studies demonstrate that FXYD5 increased the apparent affinity for cytoplasmic
+ Na twofold (K0.5 9.24 ± 2.68 mM for vector, 4.53 ± 1.54 for FXYD5 transfected cells), while Vmax values were not significantly changed. In contrast, FXYD5
+ induced a significant 60% decrease in the apparent affinity for K (K0.5 0.07 ±
0.01 mM for vector, 0.12 ± 0.01 for FXYD5) and increased Vmax twofold. Stable transfection of vector or FXYD5 did not affect surface expression of the Na,K-
ATPase, indicating that FXYD5 specifically altered pump kinetics. These effects could be due to either a change in sodium binding sites or the rate constants that stabilize the sodium binding conformation of the Na,K-ATPase. Since we observed a decrease in the apparent affinity and increase in Vmax for extracellular
K+, this suggests that FXYD5 increases the rate of K+ deocclusion:
E2(K)ATPE1ATP. Furthermore, Scatchard analysis indicates FXYD5 decreases the negative cooperativity observed in Na+ binding, suggesting that FXYD5 affects the E1P-E2P conformational equilibrium by shifting towards E1P [157]. The notion that the functional role of FXYD5 is to increase the catalytic efficiency of the Na,K-ATPase is reinforced by a 50-80% increase in the specificity constants of the Na,K-pump for both cations. These observations have implications for tissues such as the kidney and lung where sodium homeostasis is important for
water absorption, where cells that express FXYD5 might be expected to have a higher catalytic turnover and increased Na+ absorption (Figure 3-8).
We used both in vitro and in vivo models of increased sodium absorption to evaluate FXYD5 expression. MDCK-ENaC cells stably express the α, β and γ
ENaC subunits upon induction, and we demonstrated by immunofluoresence and
86Rb+ uptake that ENaC induction also increased Na,K-ATPase expression and activity. Interestingly, there was a significant, 75% decrease in FXYD5 expression, suggesting that sodium hyperabsorption downregulates FXYD5. This result was confirmed by quantitative RT-PCR and immunoblot analyses in tissue from the lungs of mice that specifically overexpress ENaC in airway epithelia. A similar decrease in FXYD4 (CHIF) expression is observed in mouse kidney medullary collecting duct cells during hyperkalemic conditions experienced in acute tubular necrosis [158]. In these two examples, conditions that led to a marked upregulation of Na,K-ATPase resulted in downregulation of the FXYD congener. This may represent an attempt to restore homeostasis, shifting the fine-tuning effects of FXYD5 and decreasing the catalytic efficiency in favor of higher enzyme activity.
In CF airway epithelia, there is marked increase in ENaC-mediated hyperabsorption of sodium, and early studies indicated that there was an upregulation of the Na,K-ATPase as well [42]. Based on our observations that upregulation of ENaC, either over 48 hr or chronically in a mouse model, downregulates FXYD5 expression, we expected FXYD5 expression to be downregulated in CF as well. However, in multiple CF models, FXYD5
81 expression was increased. FXYD5 was significantly increased almost 3-fold in nasal epithelia of CF mice by quantitative RT-PCR, on immunoblots from murine nasal epithelia and by immunohistochemical staining and immunoblots of lung epithelia of CF mice. Although in CF mouse lung, ENaC activation is less than is seen in the human condition, ENaC upregulation is clearly demonstrated in CF mouse nose. The findings in mice were mirrored in human nasal scrapes where
FXYD5 is upregulated in CF compared to controls. Upregulation of ENaC activity is prominent in human CF nasal epithelium. Finally, in human tracheal epithelial cells grown at the air-liquid interface, application of the CFTR inhibitor I-172 for
72 hours resulted in upregulation of FXYD5, confirming in another human model that lack of CFTR function increases FXYD5 expression. In this model, ENaC upregulation is not observed, but a CF inflammatory phenotype does occur [154].
In these cells, inhibition of NF-κB with PDTC returned FXYD5 expression to control values, supporting the notion that the upregulation of FXYD5 in CF might be entrained not by the increased sodium load but by the proinflammatory state of the airways. To further investigate this possibility, we stimulated HTE cells with the pro-inflammatory cytokines TNF-α/IL-1β and saw a significant increase in
FXYD5 mRNA expression, which was blocked by another inhibitor of NF-κB.
FXYD5 has been shown to increase cell motility and has been implicated in the regulation of pro-inflammatory cytokines such as MCP-1 [116]. Interestingly, the
MCP-1 gene has a κB site upstream of its promoter, suggesting that increased
NF-κB activity observed in CF epithelia may also contribute to the MCP-1 autoregulatory loop previously described to be affected after FXYD5 inhibition
82 and that these effects are downstream, feed-forward regulators of FXYD5 expression.
In patients with CF, a prominent hypothesis for the pathogenesis of the airway disease postulates that the airway surface liquid (ASL) liquid that lines the epithelia of the lung is dehydrated due to increased ENaC activation [36, 40], which promotes bacterial adherence, infection and inflammation. Given that our kinetic data indicate that FXYD5 increased the catalytic efficiency of the pump for
Na+ and K+, increased FXYD5 expression may contribute to further ASL dehydration by modulating Na,K-ATPase activity to increase transepithelial Na+ absorption down the ENaC/Na,K-ATPase axis (Figure 3-8). Furthermore, the inflammatory phenotype in CF may contribute to ASL dehydration by increasing
FXYD5 expression, as well as being exacerbated by it. We conclude that FXYD5 modulates Na,K-ATPase pump activity, increasing transepithelial Na+ absorption, and suggest that increased FXYD5 expression observed in CF airway epithelia may therefore contribute to airway surface dehydration.
Acknowledgements
We gratefully acknowledge Alma Wilson, Christian VanHeeckeren, Veronica
Peck and the Cystic Fibrosis Animal Core for their technical assistance. We also thank Dr. Thomas Kelley for his assistance in preparing mouse nasal epithelium and Yongyi Qian for her help with immunohistochemistry. This work was supported by National Institute of Health grants P30DK27651 and T32HL07418 and from the Cystic Fibrosis Foundation. Pamela B. Davis gratefully
83 acknowledges support from the Arline and Curtis Garvin Research
Professorship.
84
Figure 3-1. MDCK-hF5Flag cells stably express FXYD5-Flag at cell membrane. MDCK cells were stably transfected with pThF5kFlag or sham vector control and stained with anti-Flag antibody coupled to anti-rabbit Alexa-fluor 568
(red, A,C) and anti-α1-Na,K-ATPase antibody coupled to anti-mouse Alexa-fluor
488 (green, D,F) antibodies. Nuclei were counterstained with Hoechst dye (blue).
Colocalization of FXYD5-Flag and Na,K-ATPase is observed (merge, E) compared to sham control (merge, B). Immunoblot analysis of FXYD5-Flag, α1-
Na,K-ATPase or actin loading controls demonstrating expression of FXYD5-Flag, but unchanged expression of Na,K-ATPase between sham and FXYD5-Flag cell lines (G). (H) Densitometric analysis of 4 blots of α1-Na,K-ATPase membrane preparations shown in (G) normalized to actin, demonstrating no significant change in α1-Na,K-ATPase.
Figure 3-1
A B C
D E F
hF5-Flag merge α1 Na,K-ATPase
H G Sham hF5Flag NS
-Na,K- α1 (4) (4) ATPase
Flag
Actin
Figure 3-2. FXYD5 modulates Na,K-ATPase activity. MDCK-hF5Flag (close triangles) or MDCK-sham control (closed squares) cells were preincubated 15 min. with or without 100 µM ouabain as described under Experimental
Procedures. 86Rb+ fluxes were measured over 6 minutes at (A) constant external
NaCl and varying external RbCl (0-2000 µM) or (B) constant external RbCl and varying external NaCl (0-70 mM). Inset, Scatchard analysis indicating (inset A) non-cooperativity of Rb+ binding or (inset B) negative cooperativity of Na+ binding. Initial Ouabain-sensitive 86Rb+ (K+) influx rates were fit to Equation 1
(non-cooperative model) and represent 6 independent experiments ± S.E. Kinetic constants are summarized in Table I.
Figure 3-2 A
uptake + Rb 86 (pmol/mg*min)
B
uptake + l/mg*min) Rb 86 (pmo
Kinetic constants K+ Na+ Specificity Specificity Model Cell line K0.5 Vmax constant K0.5 Vmax constant mM mM
Non-cooperative (Equation 1) Sham 0.153 6512 1 2.866 10905 1 hF5Flag 0.205 12735 1.44 1.648 9780 1.56
Cooperative (Equation 2) Sham 0.07 5356 1 9.24 8858 1 hF5Flag 0.12 10240 1.12 4.53 7956 1.83
Table 3-1. Comparison of intracellular Na+ and extracellular K+ activation of
Na,K-ATPase pump activity in FXYD5 transfected MDCK cells.
Values for apparent kinetic constants of human FXYD5 transfected MDCK or sham control cells were calculated using the non-cooperative and cooperative models of ligand binding (Equations 1 and 2, respectively, under Experimental
Procedures). Specificity constants were calculated according to Equation 3.
Values shown are the averages ± S.E. of 6 independent experiments.
Figure 3-3. ENaC activation increases Na,K-ATPase expression and activity but downregulates FXYD5. MDCK-ENaC cells were incubated for 48 hours in serum-free media (control, A,C) or induction media (induced, B,D) to stimulate the expression of the epithelial sodium channel (ENaC). Cells were incubated with antibodies against the α1-subunit of the Na,K-ATPase, stained using anti- mouse IgG antibodies labeled with Alexa-fluor 568 (red) and imaged for 150 ms.
Nuclei were counterstained with Hoechst dye (blue). Na,K-ATPase activity was assessed by 86Rb+ uptake (E) and demonstrated a 6-fold increase in MDCK-
ENaC induced cells compared to controls (n = 10, * = P<0.0001). Quantitative
RT-PCR analysis revealed a 75% decrease in FXYD5 expression in induced vs. control cells (F). Data were normalized to GAPDH expression (n =6, * =
P<0.0001).
Figure 3-3
Control Induced
A B
C D
E F
*
*
Figure 3-4. FXYD5 is decreased in lungs from mice that overexpress the
Scnn1b transgene. Transgenic mice overexpressing the Scnn1b (ENaC β- subunit) transgene were assessed for FXYD5 expression. (A), Quantitative RT-
PCR analysis of lung tissue from ENaC +/- mice exhibited a significant, 40% decrease in Fxyd5 expression (n = 4, * = P<0.0001) compared to wild-type control littermates. Data were normalized to Gapdh expression. (B),
Representative immunoblot analysis of crude membrane preparations of ENaC
+/- vs. -/- lung tissue. The immunoblot was cut in half, stained with either FXYD5 antibody 562 or actin as a loading control and demonstrates a decrease in
FXYD5 expression in ENaC +/- mouse lung.
Figure 3-4
A
*
B
Figure 3-5. FXYD5 is upregulated in the nasal epithelia of S489X-/- CF mice.
(A) Quantitative RT-PCR analysis revealed that FXYD5, normalized to Gapdh, was increased in nasal epithelia from S489X-/- CF mice compared to S489X +/+ littermate controls (+/+, n=4; -/-, n=5; * = P<0.001). (B) Immunoblot, representative of 4 independent experiments, of membrane preparations of nasal epithelium from S489X+/+ and S489-/- mice using FXYD5 562 antisera and actin loading control.
Figure 3-5
A
*
B +/+ -/- FXYD5 Actin
Figure 3-6. FXYD5 is increased in airway epithelia of CF mice. Upper airway and lung tissue was collected from adult ΔF508-/- mice (B,D,F) and their wild type littermate controls (A,C,E). A-D are representative images taken from upper airway (A,B) or lung (C,D) and stained with affinity purified 562 FXYD5 antibody.
Arrows denote positive staining airway and alveolar epithelial cells. (E,F) are representative of no-primary antibody upper airway controls. Images taken under
40x objective lens. (G) Representative immunoblot demonstrating increased
FXYD5 expression in crude membrane preparations from ΔF508 CF mice compared to wild type control littermates.
Figure 3-6
Wt ΔF508/ΔF508 (CF) A B . .
Upper Airway
C D
Lower Airway
E F
. . No Primary Control
G . Wt CF
FXYD5
Actin
Figure 3-7. CFTR inhibition upregulates FXYD5 in human airway epithelia.
(A) RNA was prepared from 4 CF and 7 non-CF human nasal scrapings as described in Experimental Procedures. Parallel samples were used to normalize copy number according to % epithelial cells retrieved in scrapings, which varied from 84-95% or 77-98% in CF or non-CF samples, respectively (* = P<0.02). (B)
Quantitative RT-PCR analysis of FXYD5 in human tracheal epithelial cells treated with CFTRinh-172 for 3-days increased FXYD5 expression, which was abrogated by including NF-κB inhibitor PDTC (control, n=11; inh-172, n=7;
PDTC, n=4), * = P<0.001 vs. control, # = P<0.01 vs. Inhib-172. (C) Quantitative
RT-PCR analysis of FXYD5 in HTE cells 4 hours after stimulation with TNF-α/IL-
1β. Increased FXYD5 expression after TNF-α/IL-1β stimulation was inhibited by pretreatment with the NF-kB inhibitor BAY 11-7085 (control, n=10; TNF-α/IL-1β, n=9; TNF-α/IL-1β + Bay 11-7085, n=6), * = P<0.001 vs. control, # = P<0.01 vs.
TNF-α/IL-1β. Data were normalized to GAPDH and analyzed by one-way
ANOVA using Newman-Keuls post-test analysis.
Figure 3-7
A *
B *
#
C * #
Figure 3-8. Model of FXYD5 role in Na+ absorption in CF airway epithelia.
Due to the loss of CFTR mediated Cl- efflux and resulting ENaC-mediated Na+ hyperabsorption, Na,K-ATPase expression and activity is increased in CF airway epithelia. Expression of FXYD5, a regulator of the Na,K-ATPase, is also upregulated in CF epithelia and increases the rate of Na+ absorption along the
ENaC/Na,K-ATPase axis. As a result, FXYD5 contributes to the chronic dehydration of the protective airway surface liquid that lines the airway.
Figure 3-8
Airway Surface Liquid + + Na Na
ENaC CFTR ENaC CFTR
Cl- Airway epithelial + + Cell Na Na+ Na
Na,K -ATPase Na,K- FXYD5 + ATPase + K K+ K
Normal CF
Chapter 3 ADDENDUM: FXYD5 is increased in the lungs of CF mice after infection with P. aeruginosa.
Timothy J. Miller1,2, Aura Perez2, and Pamela B. Davis2 From the Departments of 1Pharmacology and 2Pediatrics, School of Medicine, Case Western Reserve University, Cleveland, Ohio
(This section includes data that are relevant to Chapter 3, but are not included in the manuscript).
Introduction
Patients with cystic fibrosis (CF) have mutations in the gene encoding the cystic fibrosis transmembrane conductance regulator (CFTR), a cAMP-activated chloride channel located in the apical membrane of most epithelial cells.
Alterations in CFTR lead to defective ion transport in many organs, such as the pancreas, liver, and intestine, although the majority of complications result from chronic inflammation and infection in the lung. Typically, CF airway epithelia exhibit increased expression of pro-inflammatory cytokines and mucus accumulation, leading to an airway environment that is predisposed to oxidative stress and inflammation. As a result, most mortality in humans results from persistent and recurring bronchopulmonary infections of trapped bacteria, such as mucoid Pseudomonas aeruginosa [159].
Although the exact link between CFTR defects and the inability to clear trapped, pathogenic bacteria remains unclear, it has been postulated that increased sodium absorption and decreased airway surface liquid are critical modulators of disease pathogenesis in the lung [33, 36, 40]. The upper airways
102 of CF mice demonstrate similar ion transport properties as human CF airway epithelia, and CF mice have a reduced ability to clear aerosolized pathogens such as Staphylococcus aureus compared to control animals [160]. However, unlike humans, mice with defective Cftr do not develop spontaneous lung infections [159, 161, 162]. To better evaluate the role of sodium absorption in airway defense, Mall and colleagues created a transgenic mouse that overexpresses the ENaC β-subunit [36]. They found that these mice had increased Na+ absorption, decreased ASL volume, increased mucus concentration, mucus adhesion to airway surfaces, neutrophilic inflammation and poor bacterial clearance, suggesting that sodium hyperabsorption may be a precursor in human CF lung disease.
The exaggerated pro-inflammatory response observed in CF airways is characterized by enhanced neutrophilic infiltration and secretion of interleukin-8
(IL-8) in the broncheoavleolar lavage fluid (BALF) [163]. Lack of functional CFTR has been associated with increased transcription of nuclear factor kappa-B (NF-
κB), which regulates the synthesis of IL-8, and epithelial cell lines derived from
CF patients have been shown to have increased NF-κB activity, suggesting that
NF-κB may be a key regulator of inflammation in CF [164]. We have previously shown that FXYD5 is highly expressed in the lung and is upregulated in human and mouse models of CF [165]. Furthermore, we have demonstrated that stimulation of human tracheal epithelial cells by the pro-inflammatory cytokines
TNFα and IL-1β upregulates FXYD5 expression, an effect blocked by inhibitors of NF-κB. We now show that Fxyd5 is expressed in airway epithelia of murine
103 bronchioles and that Fxyd5 expression is significantly increased in CF airway epithelia 3 hours after exposure to aerosolized P. aeruginosa. We suggest that
FXYD5 is increased in CF airway epithelia as a result of increased pro- inflammatory signaling and exacerbates airway surface liquid dehydration by increasing Na+ extrusion from airway epithelia.
104 Materials and Methods
Mouse strains. Breeding pairs of C57BL/6J mice were purchased from Jackson
Laboratory (Bar Harbor, ME) before use. Breeding pairs of heterozygote mice bearing the ΔF508 Cftr mutation in a mixed genetic background were a kind gift from Dr. Kirk Thomas from the University of Utah and were backcrossed into the
C57BL/6J background for at least 10 generations in the CF Animal Core Facility at CWRU before use. Cystic fibrosis mice were fed the liquid elemental diet
Peptamen (Nestle Clinical Nutrition, Deerfield, IL) after weaning. Mice were housed in static isolator units (Lab Products, Seaford, DE) on corncob bedding
(combination size; The Andersons, Maumee, OH). Light cycles were 12 hr on, 12 hr off. Cystic fibrosis mice for these strains are indicated by their Cftr mutation and are referred to as “CF mice”.
Laser Capture Microdissection (LCM). 8-week-old male mice bearing the ΔF508 mutation and WT littermates, backcrossed to the C57BL/J6 background, were inoculated with free P. aeruginosa (5x107 Cfu PA M57-15) by nasal insufflation and sacrificed 3 and 24 h after inoculation; uninfected mice served as control (0 h) from each group. Each time point was done in triplicate. Immediately upon sacrifice, lungs were snap frozen in liquid nitrogen, and kept at -80ºC until needed. LCM was performed following instructions from Arcturus Systems for
Microgenomics (Mountain View, CA). Briefly, slides (8 µM) were prepared from the frozen lungs same day of dissection per Arcturus instructions, one slide per cap. Airway epithelial cells lining the bronchioles were captured using the PixCell
105 II LCM system (Arcturus Systems for Microgenomics). Laser pulses were applied with duration of 1.2 ms and 80 mW power. Pictures were taken of the slide before and after capture, as well as, the cap at either 4X or 20X, but capture was done at 20X. Immediately after material was captured, 20 µl of extraction buffer from the PicoPure RNA isolation kit (Arcturus Systems for Microgenomics) was added to the cap. RNA was extracted using the PicoPure RNA kit (Arcturus), and a double round amplification performed using RiboAmp RNA amplification kit
(Arcturus). The Enzo BioArray HighYield RNA Transcript Labeling Kit (Enzo
Diagnostics, Farmingdale, NY) was used for the IVT reaction of the second amplification round. After checking for quality of the double amplified RNA using the 2100 Bioanalyzer (Agilent Technologies, Palo Alto, CA), RNA was fragmented following Affymetrix protocol and hybridized to Affymetrix
MOE430A2 chip. Data were extracted from the chips using 1) GeneChip
Operating Software (GCOS, Affymetrix, Santa Clara, CA) to determine detection call (if a gene was considered present or absent) and 2) Spotfire DecisionSite for
Microarray Analysis (Spotfire, Somerville, MA) using GC-RMA (Robust Multi- array Analysis as a function of GC-content) to determine gene expression levels and fold changes. GCRMA normalizes across all chips in a given set using quantile normalization producing identical distributions. This method uses median polish to estimate log expression robustly. BAMarray 2.0 (Bayesian ANOVA for microarray, J. Sunil Rao and Hemant Ishwaran, CWRU, Cleveland, OH) was used to determine statistically differentially expressed genes. BAMarray analysis is based on a special type of inferential regularization known as spike-and-slab
106 shrinkage which allows for an optimal balance between total false detections
(total number of genes falsely identified as being differentially expressed) and total false non-detections (total number of genes falsely identified as being non- differentially expressed. The complete set of GC-RMA values extracted from each array were fed into BAMarray and the 3 hr and 24 hr time points were compared against the 0H time point of each corresponding genotype separately; also each CF time point was compared against its corresponding time point in the WT mice. The list of significant genes obtained by feeding GCRMA data into
BAMarray was further filtered by the detection calls from Affymetrix. Only genes considered “Present” or “Moderately Present” at each individual time point were kept.
Isolation of crude membranes. Crude membranes were prepared as previously described [119]. Mouse tissue or cultured cells were isolated, washed with PBS, suspended in 25 mM imidazole, 1 mM EDTA, 250 mM sucrose and protease inhibitor cocktail (Sigma, St. Louis, MO) and manually homogenized with 35 strokes of a prechilled Dounce homogenizer. Nuclei, unbroken cells and mitochondria were separated by centrifuging at 6000 x g for 5 minutes. The supernatant was collected and saved, while the pellet was homogenized again and centrifuged. The two supernatants were combined and centrifuged at
125,000 x g to pellet crude microsomal membranes. The pellets were resuspended in 25 mM imidazole, 1 mM EDTA, 10 mM RbCl and stored at 4ºC.
107 Immunoblot analysis. Crude membrane (10-20 µg) were dissolved in 30 µl 2x
Laemmli buffer, incubated at 37ºC for 20 minutes, loaded onto 12% SDS-PAGE and resolved at 100 V for 1.5 hr. Proteins were transferred via semi-dry blotting onto nitrocellulose (Bio-Rad, Hercules, CA), blocked with 1% BSA and incubated with either rabbit polyclonal anti-FXYD5 562 (1:250) antibody or antibody against actin (1 mg/ml) (Sigma, St. Louis, MO) in 1% BSA/Tris-buffered saline for 1 hr at room temperature. The blot was then incubated with species specific secondary antibodies (1:4000) conjugated to HRP and visualized using Pierce’s Supersignal
Pico West detection kit.
108 Results and Discussion
The thin, protective layer of surface liquid critical for ion exchange and bacterial clearance is dramatically reduced in CF airway epithelia. To explain this observation, the preponderance of evidence favors the hypothesis that predicts isotonic sodium concentrations but decreased ASL volume due to the loss of
CFTR-mediated inhibition of the epithelial sodium channel (ENaC) and resulting sodium hyperabsorption [31]. In support of this hypothesis, Mall et. al. demonstrated that overexpression of the β-subunit of ENaC in murine airway epithelia, without any change in Cftr activity, produced the reduced ASL volume, increased sodium absorption and mucus obstruction characteristic of CF airways
[36]. Once sodium enters the apical membrane of the cell via ENaC overactivity, it must exit the cell at another site, primarily through the Na,K-ATPase, which has been reported to be increased in expression and function in CF airways.
Therefore, this system may be an important component of disease progression in
CF airways.
The Na,K-ATPase is a heteromeric enzyme, composed of a catalytic α- subunit and a heavily glycosylated β-subunit, that resides in the basolateral membrane of most epithelia. Increased expression of Na,K-ATPase subunits in vivo and in vitro has demonstrated that the Na,K-ATPase has an active role in lung liquid homeostasis [56, 166]. While FXYD2 was initially identified as the γ- subunit of the Na,K-ATPase, recently other members of the FXYD family have also emerged as tissue-specific regulators of the Na,K-ATPase [86, 96]. We have recently shown that FXYD5 modulates Na,K-ATPase activity and is
109 increased in CF airway epithelia due to increased NF-κB activity [165]. In this study, we postulate that increased Fxyd5 expression may be due to exaggerated inflammatory signaling characteristic in CF airway epithelia.
Although the CF mouse lung does not spontaneously become infected, following deliberate infection with P. aeruginosa, the pathology is similar to that observed in human CF patients [167, 168]. To test whether Fxyd5 expression is altered in response to P. aeruginosa infection in CF lungs, we analyzed lung tissue from three adult ΔF508 CF mice and three of their wild type littermate controls at 0, 3 and 24 hours post-inoculation with P. aeruginosa using laser- capture micro dissection of lung epithelia (bronchioles) and subsequent microarray analysis (Figure 3-9A-D). A representative image of a lung slice pre- laser capture (Figure 3-9A), post-capture (Figure 3-9B) and the epithelia captured (Figure 3-9C) is shown. Genechip Operating Software called “present” only Fxyd3 and Fxyd5. All other FXYD family members were called absent in these samples. Spotfire Decisionsite for Microarray analysis using GC-RMA indicated that Fxyd3 was slightly increased in control mice at 3 hours, but
BAMarray did not find this increase significant (Figure 3-9D). However, Fxyd5 exhibited a significant 5.78 fold increase in lungs from CF mice after 3 hours exposure. By 24 hours, a 2.57 fold increase in Fxyd5 expression was observed, but this was no longer significant. Similar increases were not observed in wild type mice in response to P. aeruginosa. These results implicate Fxyd5 as a putative early-response gene whose function is increased after infection in CF airways.
110
These data, taken together with our previous results demonstrating that
FXYD5 increases Na,K-ATPase activity, suggest that FXYD5 may be upregulated quickly to remove excess airway and alveolar liquid in response to this infectious and inflammatory stimulus, which probably produces exudation and transudation. This observation raises the question of whether FXYD5 is upregulated in CF epithelia by virtue of the ion transport defect or because in CF, the epithelium is often a site of inflammation. Considering the increase in FXYD5 protein in uninfected murine airways and nasal epithelium, it seems more likely that the increase is due to a smoldering, pre-inflammatory stimulus characteristic of CF epithelia. This is supported by the observation that ENaC is not activated
[154] in human tracheal epithelia after treatment with CFTR inhibitor 172 and that
FXYD5 expression is decreased in our models of sodium hyperabsorption.
Therefore, we speculate that FXYD5 may uniquely contribute to ASL dehydration by synergistically driving sodium hyperabsorption as a result of increased pro- inflammatory signaling.
111
A B C
D Wild type littermate ΔF508 -/- Fold Fold Fold Fold 0 hr 3hr change 24hr change 0 hr 3hr change 24hr change FXYD3 PPA PPP 4.594 PPA -0.231 PAP PPP -1.041 PAP 0.639 FXYD5 PPA PPP 0.565 PPP 1.117 PPP PPP 5.781 PPP 2.577
E 0 hour 3 hour 24 hour Wt CF Wt CF Wt CF
FXYD5
actin
Figure 3-9. FXYD5 is increased in the lungs of CF mice after infection with
P. aeruginosa. (A-C) Murine lung tissue before (A) and after (B) laser-capture
microdissection of epithelia (C) inoculated with P. aeruginosa. (D) Detection calls
and gene expression fold changes in control or CF (ΔF508) laser-captured lung
epithelia at 0, 3 and 24 hours post-inoculation. (E) Immunoblot analysis of
FXYD5 expression in crude membranes prepared from lung epithelia of control
or CF (ΔF508) mice at 0, 3 and 24 hours post-inoculation. Actin is shown as a
loading control.
112 Chapter 4: S163 is critical for FXYD5 modulation of wound healing in airway epithelial cells
Timothy J. Miller1,2 and Pamela B. Davis2 From the Departments of 1Pharmacology and 2Pediatrics, School of Medicine, Case Western Reserve University, Cleveland, Ohio
Abstract
The FXYD family, which contains seven members, are tissue specific regulators of the Na,K-ATPase. Increased expression of FXYD5, a cancer-cell associated membrane glycoprotein, has been associated with increased cell motility and metastatic potential. To better understand how FXYD5 may modulate cell motility, we analyzed S163, a conserved residue in all FXYD family members located in the C-terminus. Ectopic expression of human FXYD5 S163 mutants in
HEK 293 cells demonstrated that negative charge at S163 (S163D) decreased membrane localization, assessed by immunofluorescence. Co- immunoprecipitation studies revealed decreased FXYD5/Na,K-ATPase interaction for S163D compared to wild-type or S163A mutants. Since the Na,K-
ATPase is critical for establishing cell polarity and suppressing cell motility, we analyzed S163 mutants in a murine airway epithelial cell wound healing model.
Wild type murine FXYD5 overexpression increased wound healing (P<0.0001), which was further increased in S163D mutants (P<0.005). However, S163A mutants inhibited wound healing compared to wild type FXYD5 overexpression
(P<0.0001). We conclude that negative charge at S163 regulates FXYD5/Na,K-
ATPase interaction and that this interaction modulates wound healing in airway epithelia.
Introduction
The recurrent remodeling of pulmonary epithelium as a result of exposure to environmental stress, viruses, and bacteria requires that airway epithelial cells migrate to wound sites and then polarize in order to maintain epithelial integrity.
The requirement to heal lesions in the airway epithelium caused by infection and inflammation might logically result in expression and activation of proteins associated with cell motility and adhesion. While numerous factors are involved in the initiation of the healing process, depolarization of the epithelial cells along the edge of the wound constitutes an intermediate step in the reorganization of actin characteristically observed during wound healing [169]. This suggests that the activity of ion channels such as the epithelial sodium channel (ENaC) and the
Na,K-ATPase may modulate the efficiency of wound repair.
While the primary function of the Na,K-ATPase, located on the basolateral surface of most epithelia, is to exchange three intracellular sodium ions for two extracellular potassium ions, the Na,K-ATPase may also propagate external stimuli within the cell [170, 171]. In particular, signals derived from the β-subunit of the Na,K-ATPase are essential for the development of epithelial cell polarity and suppression of cell motility [66, 172-174]. The Na,K-ATPase is regulated by members of the FXYD protein family, small type-1 transmembrane proteins characterized by a signature 35-residue domain containing an invariant, extracellular PFXYD sequence [45]. Currently, the role of FXYD proteins in the regulation of Na,K-ATPase signal transduction and the effect of this association on cell motility and wound repair is unknown.
Recently, members of the FXYD family have been identified as potential markers of tumorigenesis. In particular, increased expression of FXYD5, also known as
Dysadherin, has been correlated with increased tumor progression and invasiveness [73, 107, 109]. Knockdown of FXYD5 expression has correlated with decreased cell motility, whereas transfection of FXYD5 into liver cells led to decreased cell-cell adhesion, increased cell motility and diminished expression of
E-cadherin [105, 107]. Overexpression of FXYD5 also increased cortical F-actin and membrane filopodia, two prerequisites for wound closure [105, 107], and implies that FXYD5 may be a critical determinant regulating the role of the Na,K-
ATPase in cell adherence and motility. Previous reports have shown that FXYD5 is expressed in the basal layer of squamous epithelia as well as within the lung
[45]. Therefore we investigated how a conserved serine residue affects
FXYD5/Na,K-ATPase association and how this altered cell motility in an in vitro model of epithelial wound repair.
115 Materials and Methods
Cell lines. The mouse lung epithelial cell line LA4 and human embryonic kidney
(HEK) 293 cells were obtained from the American Type Culture Collection
(Manassas, VA). LA4 cells were grown in Kaighn’s modification of F12 medium
(F12K, Mediatech, Inc., Herndon, VA) and HEK 293 cells were grown Earle’s modification of MEM media (Mediatech). All media was supplemented with 10% heat-inactivated FBS.
RT-PCR, cloning and site-directed mutagenesis of FXYD5. FXYD5 cDNA was isolated by RT-PCR using the Superscript II One-Step RT-PCR kit (Invitrogen,
Carlsbad, CA) and primers designed from accession numbers NM014164
(human) and NM008761 (mouse), which contained a HindIII and NotI restriction site on the 5’ and 3’ end respectively (appendix 2). The following RT-PCR conditions were used: reactions were incubated at 50ºC for 30 min., followed by
2 minute initial denaturation at 95ºC and 40 cycles of 94ºC, 1 min. denaturation,
1 min. 58ºC primer annealing, 45 sec. of primer extension. RT-PCR products were digested with HindIII and NotI restriction enzymes and agarose gel purified using the Qiaquick gel purification kit (Qiagen, Inc., Valencia, CA). cDNAs were subcloned in pBSK2 vector to create pBhF5k (human) and pBmF5k (murine). To generate an N-terminus Flag tag in human FXYD5 that did not alter the N- terminus signal sequence, codon Q22 was mutated using the Quickchange site- directed mutagenesis kit (Stratagene, Inc., Cedar Creek, Texas). A silent mutation was introduced (CAG mutated to CAA) to create a new restriction site,
Acl1, and an M2-Flag tag inserted to produce pBhF5kQ22Flag. To create a C-
terminal Flag-tagged FXYD5, pBhF5k was digested with NotI and TfiI, agarose gel purified and used to ligate an in-frame Flag-tag (see Appendix 2). Similarly, a
C-terminus Flag-tag was inserted into murine FXYD5 to create pBmF5kFlag. The original and modified versions of FXYD5 were then subcloned into the previously described pKCERegfpSV expression vector [152] using HindIII/Not to create pKCERhF5kFlag, pKCERhF5kQ22Flag and pKCERmF5k. The Quickchange site-directed mutagenesis kit was used to introduce alanine or aspartic acid at serines 163 to create pKCERhF5kS163A, pKCERhF5kQ22FlagS163A, pKCERhF5kS163D, pKCERhF5kQ22FlagS163D, and pKCERmF5kS163A and pKCERmF5kS163D, respectively.
Transfection of LA4 and HEK 293 cells. Cells were plated in 10 cm tissue cultures plates (Costar) at 75% confluency. After 24 hours medium was changed to serum-free Optimem (Invitrogen, Carlsbad, CA) and cells were transfected with 50 µl Lipofectamine 2000 (Invitrogen, Carlsbad, CA) and 20 µg plasmid DNA per plate. Lipid-DNA complexes were incubated 4 to 5 hours on cell monolayers at 37ºC/5% CO2 after which transfection medium was replaced with complete medium. Cells were then cultured for an additional 24-36 hours and then assayed.
Isolation of crude membranes. Crude membranes were prepared as previously described [119]. The pellets were resuspended in 25 mM imidazole, 1 mM EDTA,
10 mM RbCl and stored at 4ºC.
117 Immunoprecipitation and immunoblot analysis. Crude membrane protein (125 µg) from HEK 293 cells was resuspended in 10 mM RbCl and 200 µM ouabain and solubilized as previously described [119]. Aliquots of solubilized membrane were incubated with 1 µg antibody raised against the α1 subunit of the Na,K-ATPase
(Upstate Biotech, Inc., Charlottesville, VA), 1 µg M2 Flag antibody or non-specific
IgG at 4ºC for 3 hours. 50 µl of Protein G agarose beads were added and incubated overnight on a rocking platform at 4ºC. The beads were washed two times with 25 mM imidazole, 1mM EDTA, 100 mM RbCl, 200 µM and 0.2mg/ml
C12E10, resuspended in 50 µl laemmli buffer, incubated at 37ºC for 20 minutes and loaded onto 12% SDS-PAGE. After semi-dry transfer to nitrocellulose, proteins were blocked with 1% BSA and incubated with antibodies raised against either mouse anti-α1 subunit of the Na,K-ATPase (1 µg/ml) or anti-M2 Flag tag
(1mg/ml) (Sigma, St. Louis, MO) in 1% BSA/Tris-buffered saline for 1 hr at room temperature. The blot was then incubated with species specific secondary antibodies (1:4000) conjugated to HRP and visualized using Pierce’s Supersignal
Pico West detection kit.
Indirect immunofluorescence. HEK 293 cells were cultured in 4-well chamber slides (Permanox) to 80% confluence and transfected with 4.5 µl Lipofectamine
2000 (Invitrogen, Carlsbad, CA) and 1.5 µg DNA per well. At 24-36 hrs post- transfection, monolayers were washed once with phosphate buffered saline
(PBS), fixed with 2% paraformaldehyde/PBS for 10 minutes and washed with
PBS twice. Non-specific antibody binding was blocked by addition of 4% bovine
118 serum albumin (Invitrogen, Carlsbad, CA) for 45 minutes. Primary anti-M2 Flag mouse antibody (2mg/ml) was diluted in 1% BSA/PBS, incubated on monolayers for 45 minutes and aspirated. Monolayers were washed twice with PBS and then incubated in 1%BSA/PBS containing goat anti-mouse or goat anti-rabbit IgG
Alexa-Fluor 568 (1:250, Stratagene, Inc., Cedar Creek, Texas) for 45 minutes.
Cells were washed twice, the nuclei counterstained with Hoechst (Stratagene,
Inc., Cedar Creek, Texas), mounted with Fluormount-G, and allowed to dry overnight. Immunofluorescent localization was assessed on a Zeiss 200M
Axiovert inverted microscope, with a DG4 switchable fluorescent light source
(Sutter Instrument Company, Novato, CA) and a 12-bit CoolSnap HQ camera
(Roper Scientific, Tucson, AZ) under control of MetaMorph v 6.2 (Molecular
Devices, Sunnyvale, CA). Images were obtained with a 63X numerical aperture
1.3 fluar lens using excitation and emission filter passbands of 260 ± 20nm and
645 ±30 nm, respectively. Typical exposure times for individual frames were 200 ms.
Wound Assay. Mouse LA4 airway epithelial cells were seeded at 80% confluency in 12-well plates (Costar, Inc., Cambridge, MA) and transiently transfected as described above. Transfection efficiency was assessed using GFP fluorescence of parallel-transfected wells. 24 hours post-transfection, wells were scraped with a 200 µl yellow pipette tip and the wound was imaged. Each well was wounded three times, and the wound distance was measured at 3 points along each wound to establish baseline values. Sixteen hours later, the wounds were
119 measured again. Data were expressed as the % of the gap that had been closed during that period. Sigma Stat (Systat Software Inc., San Jose, CA) was used to calculate ANOVA statistics using the Student-Neuman-Kuels regression analysis for pairwise comparisons.
120 Results
Mutations in Ser163 alter FXYD5 cellular localization. We inserted a
Flag-tag into the N- or C-terminus of human or mouse FXYD5 and transiently transfected HEK 293 or LA4 cells (Figure 4-1). Immunoblot analysis demonstrated expression of all constructs, however the N-terminal Flag-tagged human FXYD5 appears to be more efficiently translated and detected in the membrane preparations of HEK 293 cells (Figure 4-1, lane 3) compared to the C- terminal construct (Figure 4-1, lane 2). Although location of the Flag tag did not affect the apparent molecular weight of either species (human, 37 kDa; mouse,
25 kDa), the C-terminus Flag tag is inefficient for immunofluorescent detection in vivo (data not shown), as others have shown [119]. The Q22Flag construct, however, is efficiently detected in the membrane of HEK 293 cells (Figure 4-2).
Similar results were observed in LA4 cells (data not shown). Therefore, N- terminal Q22Flag-tagged human FXYD5 was used to analyze S163 mutations in human cells.
In HEK 293 cells transiently transfected with pKCERhF5kQ22Flag, there is clearly a stronger localization of FXYD5 at homotypic cell borders suggesting that FXYD5 may have a role in cell-cell adhesion (Figure 4-2C,D). S163 is conserved across all human FXYD family members, including FXYD5, the most divergent FXYD family member. We mutated S163 to alanine or aspartic acid to either remove a potential phosphorylation site or simulate addition of negative charge, respectively, and transiently transfected HEK 293 cells with N-terminal
Flag-tagged FXYD5 vectors (Figure 4-2). Anti-Flag antibodies demonstrate
indirect immunofluorescence staining of transfected Flag tagged wild type and
S163A mutants localized to the cell membrane (Figure 4-2C-F). However, HEK
293 cells transfected with Q22Flag-tagged FXYD5 containing the S163D mutation become more spindle shaped, show less FXYD5 at the cell membrane and increased intracellular FXYD5 staining (Figure 4-2G,H). These data suggest that negative charge at S163 regulates FXYD5 membrane insertion and suggest a possible regulatory mechanism for FXYD5/Na,K-ATPase interaction that may lead to altered cell motility.
S163 mutations alter FXYD5/Na,K-ATPase interaction. Members of the FXYD family, including FXYD5, are reported to interact with the Na,K-ATPase. For
FXYD5, this offers a mechanism by which FXYD5 can regulate cell motility and adhesion, since the Na,K-ATPase is implicated in these processes. Since S163D mutations alter the cellular localization of FXYD5, we sought to determine if S163 mutations affect FXYD5/Na,K-ATPase interaction by co-immunoprecipitation analysis in HEK 293 cells. Human Q22Flag-FXYD5/Na,K-ATPase complexes were solubilized in the non-ionic detergent C12E10, immunoprecipitated with either anti-α1 Na,K-ATPase or anti-Flag antibodies and identified by Western blot analysis. As previously observed, FXYD5 interacts with the α subunit, although the efficiency of co-immunoprecipitation of the tagged FXYD5 may be less due to the presence of other FXYD proteins in HEK 293 cells (Figure 4-3A) [119, 175] which can also interact with the α subunit. Serine to alanine mutations did not alter interaction with the Na,K-ATPase. However replacing S163 with a
122 negatively charged amino acid such as aspartic acid appeared to inhibit the
FXYD5/Na,K-ATPase interaction (Figure 4-3A,B). We observed decreased total
S163D protein in crude membrane preparations, consistent with our observations with indirect immunofluorescence (Figure 4-3B). Therefore, FXYD5 interacts with
Na,K-ATPase and negative charge at S163 can disrupt this interaction.
S163 modulates wound healing in murine airway epithelia. Overexpression of FXYD5 has been shown to increase cell motility. To test whether FXYD5 effects wound healing, we transfected the LA4 murine airway epithelial cell line with murine FXYD5 or control vectors and performed a scratch wound assay to determine the effect of FXYD5 S163 mutants on wound healing. Similar to overexpression in HEK 293 cells, the amount of membrane-localized Na,K-
ATPase in LA4 cells was unchanged (data not shown). Overexpression of wild- type FXYD5 (Figure 4-4A) clearly increased wound healing compared to control
(P<0.0001). Nevertheless, S163A mutants inhibited wound healing whereas
S163D mutants accelerated it (P<0.005, Figure 4-4A,B). Wound healing results were not due to altered cell proliferation rates as measured by BRDU staining
(data not shown). These data support previous findings that increased expression of FXYD5 increases cell motility, and indicate that negative charge
(mimicking phosphorylation) at S163 modulates cell adhesion through altered
FXYD5/Na,K-ATPase interaction.
123 Discussion
Although a primary function of the Na,K-ATPase is to maintain the monovalent cation balance between the interior and the exterior of the cell, it is also a critical participant in establishing polarity of epithelial cells and in cell motility. Rajasekaran and colleagues have shown that the Na,K-ATPase β- subunit is required for mediating E-cadherin-dependent cell-cell adhesion and is able to suppress invasion of carcinoma cells, suggesting that the Na,K-ATPase has a role in cell physiology in addition to its function as an ion pump [62, 64, 66,
134, 176]. FXYD5 has been reported to have an inverse relationship with E- cadherin and its overexpression in cancer cells reduces E-cadherin at the cell surface and promotes metastasis. The mechanism by which this effect is mediated may include regulation of the Na,K-ATPase pump number or activity.
We found that FXYD5 coimmunoprecipitates with the α subunit of the Na,K-
ATPase, and in addition, transfected FLAG-FXYD5 appears in the basolateral membranes of epithelial cells, the location of the Na,K-ATPase.
Such an association may be expected. FXYD5 is a member of a small family of tissue specific regulatory subunits of the Na,K-ATPase. Found primarily in the lung, kidney, lymphocytes and squamous epithelia, FXYD5 has been shown to interact with the Na,K-ATPase and increase maximal pump activity.
Although quantity of FXYD5 is certainly important, as demonstrated by transfection studies, it may be that its functional state is equally relevant for the overall effect of FXYD5. Functional interactions between the α/β Na,K-ATPase complex and FXYD1 (PLM), FXYD2 (γ), FXYD3 (Mat-8), FXYD4 (CHIF), FXYD5
(RIC) and FXYD7 have been shown [78, 95, 177-179]. There is good evidence that the FXYD proteins lay within a groove created by the M2, M4, M6 and M9 helices of the α subunit and that multiple α/β/FXYD contact sites occur [145, 146,
180-182]. Structural studies have demonstrated for other members of the family, though not specifically for FXYD5, that extracellular segments mediate the effect of FXYD proteins on the apparent ATP affinity, while the transmembrane domain regulates the effects of FXYD on the Na,K-ATPase cation affinities [89, 145]. We focused on serine 163, which is conserved across all family members and is predicted to lie just intracellular to the transmembrane domain [45]. It has been reported that FXYD2 can be phosphorylated by PKC at this conserved serine
[180]. Mutation of S163 to alanine had no effect on transfected FXYD5 membrane localization or cell shape when compared to wild-type transfected cells, but FXYD5 S163D transfected cells demonstrated decreased membrane localization of FXYD5-Flag and a spindle cell shape, consistent with a motile phenotype. It has been suggested that phosphorylation of some FXYD proteins, such as FXYD1, FXYD2, FXYD7 and FXYD10, regulates interaction with the
Na,K-ATPase and that FXYD phosphorylation maximizes pump activity, much like the interaction of phospholamban with the sarcoplasmic reticulum Ca-
ATPase (SERCA) [183]. It is currently unclear whether phosphorylation of
FXYD5 similarly relieves Na,K-ATPase enzyme inhibition. However, we observed that the S163A mutant co-immunoprecipitated with the α-subunit, whereas the
S163D mutant displayed reduced association, suggesting that S163 phosphorylation may inhibit FXYD5/α subunit interaction. Our studies are in line
125 with previous structural data on FXYD7 that indicate S163 is located along the face of the FXYD5 transmembrane domain that contacts the M2, M4, M6 and M9 pocket within the α subunit (Figure 5-2). We suspect that, similar to FXYD2, phosphorylation at S163 may induce decoupling of FXYD5 from the α/β complex.
However, S163 is not a consensus phosphorylation site for any known kinase contained within the Kinase Prosite database. Thus our coimmunoprecipitation data may also be the result of disrupting FXYD5 transmembrane structure, which would alter FXYD5/Na,K-ATPase interaction and potentially modulate cell motility.
In support of this proposal, we observed that the S163D mutation alters the association of FXYD5 with the Na,K-ATPase, which is required for epithelial polarization and suppression of cell motility. We sought to determine if FXYD5
S163 mutations affected cell motility in an airway epithelial cell wound model. As might be predicted from studies with ectopically expressed FXYD5, overexpression of wild-type FXYD5 increased wound healing in murine airway cells after 16 hours in a scratch wounding model. Interestingly, S163A mutants inhibited wound healing, suggesting that S163A mutants, which retain the ability to interact with the Na,K-ATPase in pulldown assay, may be a “dominant negative” mutation able to lock FXYD5 into a conformation able to associate with the Na,K-ATPase in the cell membrane but which prevents downstream signaling necessary for increased motility. Conversely, the S163D mutant, which does not interact strongly with the α subunit and does not clearly localize to the membrane, increased wound healing, indicating that disruption of FXYD5/Na,K-
126 ATPase interaction may promote signals leading to increased cell motility. We speculate that phosphorylation, or addition of negative charge at S163, disrupts the interaction of FXYD5 and the Na,K-ATPase at the surface of the cell, which promotes cell motility and wound healing. Conversely, prevention of such phosphorylation may lock the enzyme at the membrane in a state non-functional with respect to motility.
In the absence of such phosphorylation, disrupting the association of
FXYD5 with the Na,K-ATPase by altering the structure of the FXYD5 transmembrane domain may also propagate signals initiated by the Na,K-
ATPase, muck like blocking or locking a switch in the on or off position. Previous authors have demonstrated that in 2-D models of cell wounding, cell migration is the first step in repairing the epithelial monolayer. Since decreased expression of
α/β Na,K-ATPase subunits is also associated with epithelial-mesenchymal transition typically leading to more mobile, metastatic cancer cells, and overexpression of FXYD5 has recently been shown to down-regulate α/β expression oocytes, modulating the structure of FXYD5 may be a mechanism for regulating cell motility. In conclusion, we now show that FXYD5 modulates cell wound healing in airway epithelia and that S163 mutations probably have a profound effect on the ability of FXYD5 to interact with the Na,K-ATPase and to regulate wound repair.
127
Figure 4-1. Immunoblot of FXYD5-Flag in HEK 293 or LA4 cells. Immunoblot of crude membranes prepared from HEK 293 (10 µg, lanes 1-3) or LA4 (50 µg, lanes 4-5) cells transiently transfected with vector (lanes 1,4), pKCERhF5kFlag
(lane 2), pKCERhF5kQ22Flag (lane 3) or pKCERmF5kFlag (lane 5) probed with anti-M2 Flag antibody (1 µg/ml).
Figure 4-2. Mutations in Ser163 alter FXYD5 membrane localization in HEK
293 cells. HEK 293 cells transfected with control vector (A,B), pKCERhF5kQ22Flag (C,D), pKCERhF5kQ22FlagS163A (E,F) or pKCERhF5kQ22FlagS163D (G,H) were fixed in paraformaldehyde and incubated with anti-M2 Flag antibodies to detect the Flag-tag in the external N-terminus.
Nuclei were stained with Hoechst dye. Representative images demonstrate that
S163A mutations do not affect FXYD5 membrane localization whereas S163D mutants exhibit decreased surface staining and altered cell morphology.
Figure 4-2
A B
Control
C D Human Q22Flag FXYD5 wt
E F
Human Q22Flag FXYD5S163A
G H
Human Q22Flag FXYD5S163D
Figure 4-3. Ser163 modulates FXYD5 interaction with Na,K-ATPase. Crude membrane preparations from HEK 293 cells transiently transfected as described above were isolated and solubilized in C12E10 detergent. Immune complexes were precipitated using antibodies against the α-subunit of the Na,K-ATPase
(IP:α1, 3A), against the M2-Flag epitope (IP:Flag, 3B), or a non-specific control antibody (IP:con). 5% of total crude membrane input used for immunoprecipitation or immunopellet complexes were separated by SDS-PAGE and transferred to nitrocellulose. The membranes were cut in half to detect α1
Na,K-ATPase or FXYD5-Flag proteins and probed separately using either anti-
Flag antibody directly conjugated to HRP (IB:FXYD5-Flag) or anti- α-subunit
Na,K-ATPase antibody (IB:α1-NKA).
Figure 4-3
A
Vector con Wt FXYD5 S163A S163D
IB: α1-NKA
IB: FXYD5-Flag
Total IP:α1NKA Total IP: α1NKA IP:Con Total IP: α1NKA Total IP: α1NKA
B
Vector con Wt FXYD5 S163A S163D
IB: α1-NKA
IB: FXYD5-Flag
Total IP:Flag Total IP:Flag IP:Con Total IP:Flag Total IP:Flag
Figure 4-4. FXYD5 modulates wound repair in murine airway epithelial cells.
(A) Representative images of murine LA4 airway epithelial cells transfected with control vector, pKCERmF5k, pKCERmF5kS163A or pKCERmF5kS163D immediately following or 16 hours post-wounding. (B) Wound healing was significantly increased after transfection with wild type FXYD5 (n=8, *=P<0.0001).
S163A mutations inhibited wound healing, whereas S163D mutations increased wound healing (** = P<0.005 vs wt).
Figure 4-4
A
Control Wt S163A S163D
0 hr
16hr
B
**
*
** *
CHAPTER 5: SUMMARY, CONCLUSIONS AND FUTURE DIRECTIONS
Summary
Ever since the 19th century observation that frog skin separating two saline solutions exhibited a spontaneous electric potential difference, the concept of transepithelial salt transport has driven the philosophical and empirical quest to understand the origin of life [184]. It wasn’t until 1951, when Hans Ussing showed that active sodium transport was the source of electric current in isolated frog skin that vectoral ion transport in the absence of an electrochemical potential gradient was believed possible [185]. It was soon appreciated that ion fluxes could be coupled to cellular metabolism and thus drive the net transport of sodium across a cell [186, 187]. The Na,K-ATPase, or sodium pump, then became the star player in the field of membrane transport when it was found that the gradient originated by the Na,K-ATPase could also propel the unidirectional transport of amino acids, sugars and numerous other ions via co- and countertransporters.
Interest in the Na,K-ATPase was rejuvenated in the late 1990s with the observation that the hitherto uncharacterized accessory protein, the gamma subunit, was able to induce cation channel activity in Xenopus oocytes and stabilize the E1 conformation of the enzyme [82, 97, 188]. These studies were the first steps in the identification of the FXYD family, which was formally classified in 2000 by Sweadner et al. and has led to the subsequent
135 characterization of 10 family members [189]. The exact function of FXYD5, structurally the most divergent member of this family, has remained elusive, although roles in oncogenesis and ion transport have been suggested. The goal of this dissertation was to elucidate the effects of FXYD5 on Na,K-ATPase activity and determine the role of FXYD5 in cystic fibrosis airway epithelia. This chapter reviews the significance of the findings, highlights unanswered questions and suggests future studies.
FXYD5 modulates Na,K-ATPase pump affinity for Na+ and K+
Compelling evidence has been collected over the past several years to show that FXYD family members specifically associate with the α/β complex and modulate pump affinity for Na+, K+ and/or ATP. As has been demonstrated in numerous expression systems, Na,K-ATPase activity is not dependent on FXYD association but rather is enhanced or decreased by interaction with FXYD proteins. Using a well-characterized epithelial cell line that exhibited low-levels of endogenous FXYD5 expression, we developed a model system to examine the effects of FXYD5 overexpression on Na,K-ATPase kinetics. We found that
FXYD5 significantly increased the K0.5(K+) by approximately 60% and increased the K+ dependent maximal Na,K-pump uptake 2-fold. In contrast, FXYD5
+ decreased the K0.5(Na+) 2-fold without significantly affecting the Na dependent maximum pump rate. The specificity constant, Vmax/K0.5, was calculated and used to compare Na,K-ATPase pump activity in the presence or absence of FXYD5
(Table 3-1), and indicates that FXYD5 increases the overall pump efficiency for
136 both Na+ and K+. These results indicate that, similar to FXYD4, FXYD5 increases
Na,K-pump transport activity approximately 4 fold at physiological intracellular
Na+ concentrations by increasing the apparent affinity for Na+, but unlike FXYD4 also increases the maximal rate of K+ transport [78]. In CF airway epithelia,
FXYD5 would therefore function to maintain sodium homeostasis, enabling the
Na,K-ATPase to approach maximal velocity more quickly in response to increased intracellular sodium concentration.
While the expression of many FXYD proteins is tissue-localized, an increasing number of studies have shown that FXYD5 is broadly expressed in active ion transport tissues. We focused on the function of FXYD5 as a regulator of Na,K-ATPase activity as it pertains to lung function, specifically as a potential modulator of transepithelial sodium absorption in airway epithelia. As a physiological consequence of FXYD5 expression, an increase in Na+ affinity should be associated with a lower steady-state level of intracellular Na+, which can then allow the pump to efficiently respond to increases in Na+ concentration, at lower set-point levels of cytosolic Na+, and rapidly restore intracellular Na+ equilibrium. In polarized airway epithelia, this may be a revelant mechanism for maintaining intracellular sodium homeostasis, which is driven by ENaC-mediated sodium absorption through the apical membrane, which serves to regulate ASL volume. FXYD5 would therefore lower intracellular sodium concentration and help maintain the driving force for sodium reabsorption across the lumen under normal conditions. Our data indicate that under conditions of acute stimulation of apical sodium absorption via ENaC induction, FXYD5 expression and membrane
137 localization is decreased, possibly due to negative feedback signals that are as yet undetermined. However, this may be expected, as stimuli that increase
ENaC-driven sodium absorption coordinately increase Na,K-ATPase membrane translocation for increased transepithelial transport, reducing the need for fine- tuning control in favor of mass transport.
Interestingly, this is reflected in the expression pattern of FXYD5 in the kidney. In the cortical collecting duct, the half-maximal concentration for activation of the Na,K-ATPase by sodium is less than 10 mM, indicating that
FXYD5 could effectively alter Na,K-ATPase activity in response to small changes in intracellular sodium concentration. Scatchard analysis indicated that FXYD5 decreased the negative cooperativity observed in Na+ binding, which suggests that FXYD5 affects the E1P-E2P conformational equilibrium by shifting it towards
+ E1P. This may be due to decreased competition by K at cytoplasmic binding sites for Na+ or by increasing the affinity of the enzyme for ATP at the low-affinity site in the E1P conformation. FXYD5 has been observed in the cells of the collecting tubes and ducts, specifically in the intercalated cells, suggesting that
FXYD5 function or expression may be responsive to changes in the pH of the interstitial milieu or bicarbonate secretion.
We observed a decrease in the apparent affinity and an increase in Vmax for extracellular K+, which suggests that FXYD5 increases the rate of K+
+ deocclusion: E2(K)ATPE1ATP. Since ATP binding induces K release into the cytosol, FXYD5 – α/β interaction may enhance this rate-limiting step in pump activity, which may be reflected by the increase in maximal pump rate. Further
138 studies examining the effects of FXYD5 on ATP binding or enzyme phosphorylation are needed, and which α/β isoforms are affected identified.
While most FXYD protein studies are carried out in heterologous expression systems, it is unclear whether FXYD5 preferentially associates with specific α/β isoforms. Such studies would increase our appreciation of the tissue- and cell- type specific regulation of Na,K-ATPase activity by FXYD proteins. Most especially, this information would be invaluable to estimate the contribution of particular α/β isoforms to the clearance of fluid from the airways.
Potential consequences of increased FXYD5 expression in airway epithelia
The importance of lung liquid transport is manifest in the pathology of airway and alveolar diseases such as cystic fibrosis and pulmonary edema. The proper maintenance of lung liquid volume requires a balance of active ion absorption and secretion across airway and alveolar epithelium. The results of several studies have shown the coordinated removal of fluid from the distal airspace is maintained by a balance of chloride secretion and sodium absorption.
The general model for transepithelial fluid movement is that active sodium absorption drives osmotic water absorption, which is a critical factor in resolving pulmonary edema. This model has been demonstrated using pharmacological inhibitors of the apically located epithelial sodium channel (ENaC), which have been shown to reduce the amount of fluid absorption from dog, sheep and human lungs [190-193]. Similarly, inhibition of chloride secretion in cystic fibrosis airway epithelia or by chloride channel inhibitors such as glibenclamide has
139 demonstrated that the maintenance of airway surface liquid volume is important in ensuring proper depth of the periciliary fluid, which is necessary for efficient mucociliary clearance [23, 25, 194, 195]. Understanding the regulation of fluid reabsorption has led us and others to focus on the contribution of the Na,K-
ATPase in epithelial models of disordered sodium transport, such as CF [165].
Expression and activity of the Na,K-ATPase is upregulated in normal or injured airway tissue by β-adrenergic agonists, dopamine, epidermal growth factor, keratinocyte growth factor, thyroid hormone and dexamethasone to increase sodium absorption and augment lung liquid clearance [59, 60, 196-200].
Inhibition of Na,K-ATPase function with ouabain has been shown to block sodium transport in alveolar epithelial cells and edema clearance in isolated rat lungs
[57, 201, 202]. In contrast, gene transfer of the α- and β-subunits of the Na,K-
ATPase in vitro and in vivo has been shown to increase Na,K-ATPase activity and increase the rate of fluid transfer across alveolar epithelium [56, 57, 61].
These studies suggest that Na,K-ATPase subunits are critical for the vectoral transport of sodium down the ENaC/Na,K-ATPase axis in the airways.
Previous studies have shown a 2-3 fold increase in 3H-ouabain binding in membranes of CF airway epithelia from human patients, indicating an increased amount of active Na,K-ATPase [149]. We found that FXYD5 mRNA was significantly increased in nasal scrapes from human CF patients and CF knockout mice and was upregulated in HTE cells treated with a CFTR inhibitor. In parallel, we found that the protein is also increased in the nasal epithelia and lungs of CF knockout mice. Based on our findings that FXYD5 is normally
140 downregulated in models of sodium hypersabsorption, this counterintuitive discovery implicates FXYD5 as a potential modifier of disease severity in CF by contributing to the dehydration of ASL volume (Figure 5-1).
Active Na+ transport through the Na,K-ATPase is critical for maintaining dry airways, however the increased pump activity observed in CF as a result of increased FXYD5 expression may contribute to the airway dehydration and mucus accumulation observed in CF airways. The hyperabsorption of sodium across the airway epithelia is mainly attributed to increased activity and expression of ENaC. However, sodium must travel down the ENaC/Na,K-
ATPase axis and exit the cell through the basolateral membrane. Increasing evidence indicates that alterations in Na+ transport are the driving force in CF airway disease, and inhibition of ENaC activity has been proposed to ameliorate the airway pathobiology in CF. However, regulation of in vivo Na+ uptake in CF airways using aerosolized amiloride has met with only limited success due to low potentcy and rapid absorption by the airway epithelia [194, 203, 204]. Recently, a a new ENaC inhibitor has been synthesized that is 60-100 fold more potent, 2-5 fold less reversible, slower at crossing the epithelium and has 170-fold lower Koff compared to amiloride [194]. When tested in sheep, this compound increased mucociliary clearance and increased ASL volume expansion, emphasizing the contribution of sodium absorption in airway dehydration.
We speculate that FXYD5, has been shown to modulate Na,K-
ATPase activity in airway epithelial cells, might also be a therapeutic target.
Increased expression of FXYD5 in the airway epithelia of CF mice suggest a
141
Aldosterone Cystic fibrosis Diet Cancer COPD
+ Na+ Na Na+ NF-kB ENaC ASL activation
Increased + + Na uptake Inflammation Na + + Na Na Na+ Na+ + + + K K+ K K K+
FXYD5 Na,K-ATPase
+ + + Na+ Na Na Na
Figure 5-1. Regulation and role of FXYD5 expression in ASL dehydration.
Increased sodium absorption, often mediated by ENaC, causes downregulation of FXYD5 expression by an unknown mechanism(s) in the absence of inflammatory stimuli. However, pro-inflammatory stimulation in leads to an increase in FXYD5 expression, leading to conditions such as decreased airway surface liquid (ASL) volume or increased motility.
142 potential target-rich environment, which is supported by our in vitro studies demonstrating that Flag-FXYD5 is membrane localized. While complete inhibition of the Na,K-ATPase is incompatible with life, FXYD5 regulates the Na,K-ATPase in a positive manner, and this increased activity may be enhanced at the site of phosphorylatable serines. An increased understanding of the mechanism by which FXYD5 induces increased Na,K-ATPase activity, whether by simple interaction or by a more complicated signaling event, would require greater knowledge of the contribution of particular intracellular serine residues, such as
S163 and S170. Similiarly, inhibiting FXYD5 in airway epithelia might reduce the excess activity of the Na,K-ATPase while leaving intact its essential cellular functions and slow Na+ absorption along the ENaC/Na,K-ATPase axis. Thus,
FXYD5 may be a viable therapeutic target for reducing Na+ hyperabsorption and increasing mucus hydration in CF airway epithelia.
Secondary signaling effects of FXYD5/Na,K-ATPase interaction
FXYD5, also known as dysadherin, has been implicated in the metastatic progression of tumor cells in a variety of epithelial and non-epithelial derived cancers, and has been correlated with a poor prognosis for patients with tumors exhibiting increased FXYD5 expression. In some tumor models, FXYD5 appears to have an inverse correlation with E-cadherin expression, and reduction of E- cadherin is thought to be a necessary step in cancer invasion and metastasis.
Similarly, increased FXYD5 expression has been linked with increased cell motility and focal contact formation, whereas downregulation hasbeen associated
143 with enlarged, flattened cells and increased transverse actin fibers. This suggests that FXYD5 may modulate cell motility by altering cytoskeletal elements, which is further supported by the observation that pancreatic cells treated with FXYD5 siRNA have an increased amount of paxillin containing focal adhesions [107].
However, although these observations indicate a link between FXYD5 expression and tumor progression, no clear cause responsible for this upregulation has been elucidated and in fact, few clinical studies correlating E- cadherin aberrations and FXYD5 expression exist [117].
However, the maintenance of cell-cell adhesion due to E-cadherin mediated tight junction formation and the role of the Na,K-ATPase in epithelial cell polarization suggest that a functional synergism exists in the regulation of epithelial cell structure. Loss of this relationship or aberration of either E-cadherin or Na,K-ATPase function might alter cell polarity and result in epithelial- mesenchymal transition commonly observed in carcinomas [134]. Decreased expression of the β-subunit Na,K-ATPase has been observed in prostate cancer and overexpression has been shown to decrease cell motility and invasion, suggesting that in normal epithelial cells, the pump may act as a suppressor of motility. In contrast, overexpression of FXYD5 has previously been shown to increase cell motility, whereas inhibition by siRNA increases cell adhesion.
Interestingly, another argument for the physiological relevance of the effect of
FXYD5 on the affinity of the Na,K-ATPase for Na+ is suggested by the observation that, unlike FXYD2, the effects of FXYD5 on cell motility were not due to changes in proliferation rates [205, 206]. NRK-52E kidney cells, in which
144 FXYD2 was overexpressed, exhibited reduced proliferation rates, but only if the
FXYD2 variant reduced the Na+ affinity of the Na,K-ATPase, suggesting a relationship may exist between intracellular sodium concentration, growth rate and Na,K-ATPase function.
While a plethora of data has been gathered demonstrating the interaction of multiple FXYD proteins with the Na,K-ATPase α/β complex, few structural studies have investigated how FXYD proteins commute their functional effects or identified specific interacting sites. A systematic analysis studying the effects of swapping the transmembrane domains of FXYD2 and FXYD4 demonstrated that the transmembrane domain mediates the efficiency of FXYD/Na,K-ATPase association and may determine the sodium affinity of the pump [88]. A study by
Li et al. demonstrated that the transmembrane domain of FXYD7 exhibited two
“faces”, one of which appeared to be the interface with the α-subunit and modulated the effects on Na+ affinity [145]. Based on these studies, we hypothesized that Ser163 in the cytoplasmic domain adjacent to the transmembrane helix, which is conserved in all FXYD proteins, might alter
FXYD5/Na,K-ATPase interaction or function (Figure 5-2). We found that mutation of S163 to alanine maintained FXYD5/Na,K-ATPase association, whereas mutation to aspartic acid blocked FXYD5 interaction with the α-subunit and inhibited membrane insertion. The helical wheel plot of FXYD5 indicates similar presentation of conserved residues in Region A when compared with FXYD2, 4 and 7, and would suggest that these residues are necessary for stable α-subunit association. However, the residues in Region B, including S163, have been
145
V I S
T L 152 163 159 156 V Region A 148 Region B I 149 155 160 I FXYD5 TM T 162 153 L 151 A 146 158 157 G I 147 154 150 161 G L F L A
Figure 5-2. Helical wheel plot of FXYD5 transmembrane domain. Residues
146-163 are plotted similar to Li et. al., demonstrating the residues localized in two faces. Region A (pale orange), where none of the residues interact with the
Na,K-ATPase, and Region B (blue), where residues are predicted to interact with and/or modulate the Na,K-ATPase. Blue residues: isoleucine, valine, leucine; red residues: phenylalanine, threonine; green residues: conserved glycines; yellow residue, Ser163.
146 found to moderate Na,K-ATPase affinity for sodium by interfacing with the FXYD binding pocket, composed of TM2, TM4, TM6 and TM9 of the α-subunit [181,
182, 207, 208]. Interestingly, similar results were found for phospholamban, which interacts with TM2, TM4, TM6 and TM9 of the sarcoplasmic reticulum Ca+2
ATPase (SERCA) [183]. FXYD5 is unique in this regard, as residues 143, 144 and 145 are basic, unlike the rest of the family, and may shift the transmembrane helix to include S163. While our results indicate that negative charge at S163 disrupts association with the Na,K-ATPase, it is possible that we are merely disrupting a structural interaction and further studies utilizing a Ser163Pro or
Ser163Gly mutation might answer this question.
Alteration or disruption of the FXYD5/Na,K-ATPase association modulated the rate of wound healing in airway epithelia, suggesting that interaction with
FXYD proteins may also confer secondary signals to the cell. Much like the inhibition with low levels of ouabain, membrane bound FXYD5 may convey information from the extracellular milieu to regulate cell motility or adhesion. This may partially explain the relationship of FXYD5 overexpression and increased cell motility/metastasis. Chifflet et al. recently proposed that depolarization at the leading edge of epithelial cells is necessary to stimulate lamellar crawling of cells in an epithelial scratch wound model similar to ours [169]. Given the two-fold
+ increase in K dependent maximal uptake (Vmax) exhibited by FXYD5 on Na,K-
ATPase activity, FXYD5 might be functioning to restore membrane potential following depolarization of the plasma membrane potential. This may also explain the changes in actin reorganization observed after manipulating FXYD5
147 expression in pancreatic cells [107]. In the CF airway, this may be a beneficial side effect of increased FXYD5 expression as a result of the ever-present need to replenish epithelia denuded during bacterial infection.
Pro-inflammatory signals upregulate FXYD5 expression
It is been proposed that lack of functional CFTR leads to abnormal function of the NF-κB pathway. Evidence of increased NF-κB activity is supported by the presence of increased pro-inflammatory mediators such as IL-8 and IL-6 in cultures of human tracheal epithelial cells treated with a CFTR inhibitor and in BAL fluid from the lungs of CF patients. IL-10 secretion, a potent anti-inflammation chemokine which increases expression IkappaB kinases necessary for NF-κB activation, is decreased in BAL fluid from CF patients [163].
Increased nuclear localization of NF-κB has also been observed in CF epithelial cells, and exaggerated NF-κB activity is observed in CF epithelia compared to controls after exposure to bacterial challenge [209]. Strikingly, elevated chemokine and pro-inflammatory cytokine production is present in the lungs of
CF patients at birth, in the absence of bacteria, reflecting the predisposition of CF airway epithelia to inflammatory lung disease.
Chemokine expression and regulation are emerging as important factors in tumorigenesis. Many tumors express one or more chemokines that increase proliferation rates or block apoptosis, leading to autocrine and/or paracrine support of tumor growth. Cases where this exists include the overexpression of
CCL5 in prostate cancer and CXCL12 in breast cancer [116, 210]. Recently, chemokine (C-C motif) ligand 2 (CCL2) was identified as the transcript most
148 affected by FXYD5 knockdown in a breast cancer cell line [116]. FXYD5 was shown to regulate CCL2 expression in part through activation of the NF-κB pathway, and the ability of FXYD5 to promote tumor cell invasion was dependent on the establishment of a CCL2 autocrine loop. CCL2, also known as monocyte chemoattractant protein-1 (MCP-1), is a pro-inflammatory cytokine secreted from epithelial cells that is increased after airway infection to recruit and activate monocytes and basophils. Induction of CCL2 expression in keratinocytes, which prominently express FXYD5, has been linked to exaggerated inflammation in skin disorders such as psoriasis and atopic dermatitis [211]. In contrast, inhibition of NF-κB activity has been shown to decrease CCL2 secretion and attenuate inflammation.
These observations suggest a mechanism for regulating FXYD5 expression. Our results demonstrate that FXYD5 expression is upregulated in human tracheal epithelial cells in culture after stimulation with pro-inflammatory cytokines and support the notion that NF-κB activation is sufficient for FXYD5 upregulation. While we did not study CCL2 secretion in our overexpression system, it would be informative to determine if CCL2/CCR2 expression is altered or if inhibition attenuates FXYD5 mediated effects on Na,K-ATPase activity.
Interestingly, the binding pocket of chemokine (C-C motif) receptor 2 (CCR2), the receptor for CCL2, has a domain very similar to part of the extracellular domain of FXYD5 and is required for efficient ligand-receptor interaction [212]:
CCR2 - FFDYDY FXYD5 - PFFYDE
149 It would be fascinating if FXYD5 were able to act as a decoy receptor for CCL2 or if CCL2 binding to FXYD5 fostered the propagation of external signals through the Na,K-ATPase.
Endotoxemia and bacteremia increase lung vascular permeability, which can cause fluid accumulation in lung (edema). We found that FXYD5 was increased in the lungs of CF mice after challenge with P. aeruginosa compared to wild type littermates. We believe that the primary cause for this upregulation is due to the enhanced proinflammatory nature of CF airway epithelia, and suspect that FXYD5 expression is also increased in the lungs of normal mice after infection, albeit to a lesser extent. We observed an acute increase in FXYD5 transcription after apical volume expansion in cultures of human tracheal epithelial cells (data not shown), suggesting that bacterial challenge, which causes transudation of fluid as well as upregulation of NF-κB, in conjunction with fluid addition may stimulate the cell to upregulate ion transporters to drive water absorption. This may also explain how NF-κB activation, and subsequent FXYD5 upregulation, predominates over increased ENaC activity alone, which led to
FXYD5 downregulation. This is reflected in the observation that few studies have identified FXYD5 as a modifier of airway disease in CF or as a gene with significantly altered expression in the uninfected state. However, the contribution of FXYD5 to disease development may be masked if FXYD5 upregulation, and thus increased sodium absorption, is dependent on other upstream disease modifiers such as NF-κB. Increased expression of FXYD5 would therefore serve two functions, augmenting fluid absorption by increasing Na,K-ATPase pump
150 efficiency and modulating epithelial repair in the airway [213, 214]. Although each individual effect may be modest, the Na,K-ATPase may utilize as much as 40-
60% of the cellular ATP under normal physiological conditions [215]. In the lower
CF airway, where hypoxia during bacterial insult is common, Na,K-ATPase activity may comprise a large drain on cell energy reserves, exacerbating metabolic stress. Under these conditions, FXYD5 could diminish ATP usage by increasing pump efficiency and may also help maintain membrane potential by increasing K+ influx.
Future Studies
The mechanism by which FXYD5 regulates the Na,K-ATPase in airway epithelium has not been established. Others have demonstrated that murine
FXYD5 increases Na,K-ATPase activity in Xenopus oocytes and we have shown that FXYD5 alters pump affinity for Na+ and K+ in MDCK cells. Since several other FXYD family members are regulated by phosphorylation, we sought to determine if FXYD5 could be phosphorylated at S170, an intracellular serine residue. Screening for potential kinase sites revealed that S170 was a possible candidate for phosphorylation by serum- and glucocorticoid induced kinase-1
(SGK1), a cell-survival kinase originally identified in rat mammary tumors that has a high degree of similarity to protein kinases A, B and C and recognizes the consensus sequence R-X-R-X-X-S/T (Figure 5-3). Although arginine at positions
5 and 3 is necessary for efficient phosphorylation, we tested peptides containing the semi-conserved sequence K-X-R-X-X-S, corresponding to FXYD5 S170.
151 SGK1 phosphorylated human FXYD5 peptides and full-length, immunoprecipitated protein at S170, but not at S163 (Figure 5-4).
SGK1 is not absolutely necessary for Na+ homeostasis in the kidney, because SGK1-null mice compensate for reduced ENaC activity with enhanced plasma aldosterone under a standard NaCl and K+ diet, suggesting that a compensatory mechanism exists for lack of SGK1 induced ENaC activation [216-
218]. However, under conditions of low dietary sodium uptake or exposure to corticosteroids, SGK1 activity is required to increase transepithelial sodium absorption along the ENaC/Na,K-ATPase axis [217, 218]. SGK1 increases ENaC activity by reducing the rate of channel retrieval through phosphorylation of the ubiquitin protein ligase Nedd4-2, which renders it unable to interact with PY motifs on ENaC [35, 219, 220]. Recent work suggests that SGK1 is also able to directly phosphorylate an SGK1 consensus site at Ser621 in the ENaC α-subunit and stimulate ENaC activity independently of inhibition of Nedd4-2 channel retrieval [221]. SGK1 positively regulates some of the physiological effects of aldosterone on Na,K-ATPase activity by increasing membrane translocation and
α-subunit mRNA expression [222-224]. Increased expression of SGK1 has also been observed in squamous cell carcinoma and in mammary tumor formation, where overexpresssion of FXYD5 has been correlated with increased metastatic potential [225, 226]. Thus increased SGK1 expression and activity is an important contributor in the regulation of sodium transport machinery and may potentially impact FXYD5-mediated tumor formation.
152 Preliminary data suggest that phosphorylating FXYD5 S170 regulates
FXYD5 membrane insertion, similar to S163. However, we have now demonstrated that SGK1 can phosphorylate FXYD5 S170, whereas the kinase responsible for phosphorylating S163, if such exists, has not been identified.
Immunofluorescence studies of S170 mutants suggest that S170 may also regulate membrane insertion, as Flag-FXYD5 constructs containing an alanine at
S170 have decreased membrane staining and increased peri-nuclear retention
(Figure 5-5). Our previous studies have demonstrated that FXYD5 must be membrane localized to exert functional effects, thus it would be very informative to determine whether S163 and S170 mutations affect Na,K-ATPase pump kinetics. Furthermore, it would be useful to identify whether S170 is a site for protein-protein interactions or regulates ER-retrieval signals that bind to the C- terminus, which has an ER-retention signal (R-X-R).
Such an association is to be expected. Enhanced expression of SGK1 and the ENaC α-subunit have been found in model systems that recapitulate hyperglycemia associated with type 2 diabetes mellitus as well as within the kidneys of diabetic rats and humans, suggesting that increased SGK1 expression may be linked with upregulated ENaC expression [227-229]. Although alterations in SGK1 expression have not been demonstrated by gain-of-function mutations of ENaC (Liddle’s syndrome), increased cell surface ENaC density and open probability have been found in an inherited form of salt-sensitive hypertension in
Dahl rats [230]. Thus it would also be informative to examine FXYD5 expression in epithelia from the lungs and kidneys of Liddle’s syndrome patients to
153 determine if increased sodium absorption in this disorder triggers FXYD5 downregulation. SGK1 expression is also increased in type 2 alveolar cells from the lungs of CF patients compared to non-CF patients and was not due to loss of
CFTR function alone [231]. This upregulation may be particularly relevant in the disease pathogenesis characteristic of CF airway epithelia, due in part to the
ENaC-mediated hyperabsorption of sodium that is a hallmark of CF airways.
Increased transepithelial sodium absorption along the airway surface begins with entrance through ENaC in the apical membrane and ends with extrusion through the Na,K-ATPase in the basolateral membrane. SGK1 has been shown to increase Na,K-ATPase activity, much like ENaC, when coexpressed in Xenopus oocytes, an observation that has been confirmed in A6 renal epithelial cells [222-224]. While previous studies have reported that SGK1 directly activates Na,K-ATPase pump function, the exact site of SGK1 phosphorylation on a Na,K-ATPase subunit has not been reported. Our data suggest that SGK1 may regulate Na,K-ATPase activity through FXYD5, an accessory subunit which has been shown to modulate Na,K-ATPase activity
[119, 120, 165].
Our findings on the impact of phosphorylation of FXYD5 on Na,K-ATPase activity distinguish FXYD5 from other members of the family. FXYD1 (PLM) and
FXYD2 (γ subunit) can be phosphorylated by PKA and PKC, whereas phospholemman-like protein (PLMS) can be phosphorylated by PKC [177, 178,
180, 232]. The interaction of PLMS with the α/β complex has been proposed to inhibit Na,K-ATPase activity, which is relieved when PLMS is phosphorylated by
154 PKC [86, 177]. In contrast, FXYD1 appears to stimulate Na,K-ATPase activity but is inhibited upon phosphorylation. A comparison of known FXYD5 sequences
(Appendix 1) shows that S170 is highly conserved across all species sequenced to date. Our results suggest that SGK1, which can phosphorylate S170 and regulate FXYD5 membrane insertion, may play an important role in aldosterone- dependent regulation of FXYD5 in CF airways, as CF patients exhibit chronic activation of the renin-angiotensin-aldosterone (RAA) axis [38]. Based on these observations, it would also be useful to determine if FXYD5 expression or function may be regulated by exposure to mineralocorticoids and/or corticosteroids.
Our data demonstrate a relationship between FXYD5 expression, Na,K-
ATPase activity and cell motility. Taken together with the numerous studies demonstrating that FXYD5 expression is intimately linked with cell adhesion and oncogenic potential, this suggests that Na,K-ATPase expression and function may have a large impact on tumor metastases. Future studies evaluating the mechanism of FXYD5/Na,K-ATPase association must include analyses of Na,K-
ATPase subunit expression, particularly which isoforms are affected in tumor vs. adjacent tissue or in non-infected vs. infected CF airway epithelia. This will help determine how specific cell types or organs respond to altered ion transport and identify functional relationships between subunit isoforms, such as the different affinity of the α isoforms for Na+ or the role of β isoforms in cell adhesion.
The mechanism determining how FXYD5 modulates Na,K-ATPase activity is also unknown. Analyses of other FXYD proteins indicate that the
155 transmembrane domain governs interaction with the Na,K-ATPase, however
FXYD5 possesses a series of three positively charged residues at the extracellular/membrane segment interface (R143, K144, R145), which distinguishes FXYD5 from other family members. These unique residues, presented on the extracellular surface, might be utilized to scan a library of small molecules to identify potential inhibitors or activators of Na,K-ATPase activity.
This would be useful for increasing Na,K-ATPase activity during pulmonary edema or decreasing pump activity, such as in CF.
The data presented in this thesis allow speculation that altering FXYD5 expression in human tracheal epithelial cells could modulate ASL volume.
Primary airway cells are difficult to transfect with a high level of efficiency, although new technologies are being developed to increase the amount and duration of expression. Given that FXYD5 increased the catalytic efficiency of the
Na,K-ATPase, I would predict that overexpression in polarized airway cells would increase the rate and amount of ASL dehydration. In contrast, inhibition of
FXYD5 activity with short-hairpin RNA vectors could increase ASL volume expansion and protect airway cells. Proof-of-principle could be achieved by treating human tracheal epithelial cells with FXYD5 shRNA vectors and measuring ASL volume repletion/absorption after incubation with amiloride or bumetanide. In conjuction with hypertonic saline, this treatment could increase
ASL volume expansion and mucociliary clearance by slowing transepithelial sodium absorption in CF patients. Alternatively, aerosolized delivery of FXYD5 vectors could alleviate fluid accumulation in the distal airways due to pulmonary
156 edema. While each of these proposals faces significant challenges, a benefit of
FXYD5-mediated therapy is that it does not require complete inhibition or high levels of expression to mediate changes in sodium transport due to the omnipresent expression of the Na,K-ATPase. I expect that in the the near future, studies will reveal that any perturbation to cell homeostasis alters Na,K-ATPase subunit isoform expression, which will foreseeably include FXYD protein expression. If true, this will enable the targeting of specific FXYD proteins for a particular physiological state and provide opportunities for designer gene therapy. FXYD5, a modulator of Na,K-ATPase activity, may therefore be a viable therapeutic target for modifying disease severity in cystic fibrosis and cancer.
157
161 * 178 H. sapiens LTSGKCRQLSRLCRNRCR P. troglodytes LTSGKCRQLSRLCRNHCR M. mulatta LTSGKCRQLSGLCRNYCR
C. familiaris LTSGKCRQLSQFCLNRHR M. musculus LTSGKCRQLSQFCLNRHR SGK1 consensus KXRXXS
extracellular intracellular
TM Fxyd
5
Figure 5-3. FXYD5 S170 is an SGK1 consensus phosphorylation site. A comparison of FXYD5 SGK1 consensus sequences at S170, shown in red. H. sapiens, human; P. troglodytes, chimp; M. mulatta, monkey; C. familiaris, dog; M. musculus, mouse.
158 A
*
Con Wt S170A S163A
B
32 IP: P Flag
IP: IB
IB:total
Figure 5-4. SGK1 phosphorylates FXYD5 S170. (A) Peptides corresponding to residues 161-178 of the C-terminus of wt, S163A or S170A human FXYD5 were phosphorylated with SGK1. Phosphorylation of a peptide containing a conserved
SGK1 consensus site is shown as a positive control. (B) Representative immuno- or phospho-blot of HEK 293 cells transiently transfected with wt, S170A or
S170D Flag-FXYD5 vectors. Flag-immunoprecipitates were phosphorylated with
SGK1, separated by SDS-PAGE and subjected to autoradiography (top row) or immunoblot analysis against using the M2-Flag antibody (middle row).
Immunoblot analysis against the M2-Flag tag was also used to examine total protein (5 µg) content prior to immunoprecipitation (bottom row).
159 A B
WT
C D
S170A
E F
S170D
Figure 5-5. FXYD5 S170 mutations affect membrane localization in MDCK cells. MDCK cells were transiently transfected with pKCERhF5kQ22Flag, (A,B); pKCERhF5kQ22FlagS170A, (C,D); or pKCERhF5kQ22FlagS170D, (E,F). Cells were stained Hoechst dye to differentiate nuclei of untransfected cells (A,C,E) and with antibodies directed against the M2-Flag tag to assess membrane localization (B,D,E). Immunofluorescent analysis showed that the Flag antibody recognized wild-type Flag-FXYD5 in the membranes of transfected MDCK cells (A,B). The S170A mutation caused Flag-FXYD5 to be retained intracellularly, with minimal membrane staining and prominent perinuclear staining (Figure 6- 4D, arrow). In contrast, a strong immunofluorescent signal was observed in cells transfected with S170D, with little or no intracellular Flag antibody detection.
160
; 3. Canis Canis Rattus Rattus Black *************** ; ; 7. ; 11.
. Pan troglodytes Bos Bos taurus ; 2. ; 6. Echinops telfairi ; 10. Felis Felis catus Homo Homo sapiens Oryctolagus cuniculus rcellus ; 5. ; 13. ; 13. Cavia po ; 9. Tupaia belangeri ; 4. Dasypus novemcinctusDasypus Mus Mus musculus ; 12. ; rved rved FXYD domain; asterisks, membrane spanning domain. Ensemble ; 8. Appendix 1: Neighbor joinging alignment of bar, conse known FXYD5 proteins. family ENSF00000011110, by number; 1. Macaca mulatta familiaris norvegicus
1 2 3 4 5 6 7 8 9 10 11 12 13
Appendix 2
Primers for Human FXYD5 RT-PCR 5’ AAGCTTGCTAGCGCCGCCACCATGTCGCCCTCTGGTCGCCTGTGTCT 3’ AGTCGTCTAGATCACCTGCAACGATTCCGGCATAAC
Primers for Mouse FXYD5 RT-PCR 5’ AAGCTTGCTAGCGCCGCCACCATGTCACTGTCCAGTCGCCTGTGTCT 3’ AGTCGTCTAGATCACCTGTGGCGATTCAGGCAAATT
The following sequence was then inserted in frame at the Acl1 site to produce pBhF5kQ22Flag: 5’ CGGATTACAAAGATGATGATGATAAGA 3’.
The following sequence was used to create a C-terminal Flag-tag in pBhF5k to produce pBhF5kFlag: 5’ AATCGTTGCAGGGATTACAAAGATGATGATGATAAGTGATCTAGAGC
Human and mouse chimeric constructs were created by utilizing the AlwnI restriction site to produce pkCERhF5kQ22FlagmF5, pKCERhF5kmF5Flag, pKCERmF5khF5Flag.
Primers used to mutate S163: human S163A 5’ GGCATCATCATCCTCACCGCTGGCAAGTGCAGGCAGCTGTCC; human S163D 5’ GGCATCATCATCCTCACCGATGGCAAGTGCAGGCAGCTGTCC; mouse S163A 5’ GGAATTATCATTCTCACTGCTGGGAAGTGTAGGCAGTTG; mouse S163D 5’ GGAATTATCATTCTCACTGATGGGAAGTGTAGGCAGTTG
162 Appendix 3
Future directions Materials and methods.
Mouse strains. Breeding pairs of heterozygote mice bearing the sodium channel, nonvoltage-gated 1 beta, Scnn1b, under control of the rat secretoglobin, family
1A, member 1, Scgb1a1 promoter (B6;C3H-Tg(Scgb1a1-Scnn1b)6608Bouc/J, stock number 005315) were purchased from Jackson Labs (Bar Harbor, ME).
These mice are on the B6C3F1/J background and are a F1 hybrid cross of
C57BL/6J and C3H/HeJ mice. All animal studies were performed under CWRU
IACUC approved protocols.
Cell lines. Human embryonic kidney (HEK293) and Madin-Darby canine kidney
(MDCK) cells were obtained from the American Type Culture Collection
(Manassas, VA). HEK 293 cells were grown Earle’s modification of MEM media
(EMEM; Cellgro, Mediatech, Inc) and MDCK cells were grown in 1:1 (vol:vol)
DMEM/F-12 media (Mediatech). All cells were incubated in media containing
10% heat-inactivated fetal bovine serum (FBS).
Site-directed mutagenesis of FXYD5 S170. The Quickchange site-directed mutagenesis kit (Stratagene, Inc.) was used to introduce alanine or aspartic acid at serine 163 as previously described. S170 mutations were introducing using the following primers: human S170A 5’ GGCAAGTGCAGGCAGCTG
GCCCGGTTTGCCGGAATCGTTGCAGG; human S170D 5’ GGCAA
163 GTGCAGGCAGCTGGACCGGTTATGCCGGAATCGTTGCAGG to create pKCERhF5kS170A and pKCERhF5kS170D, respectively.
Transfection of HEK293 and MDCK cells. Cells were plated in 10 cm tissue cultures plates (Costar) at 75% confluency. After 24 hours medium was changed to serum-free optimem (Invitrogen, Carlsbad, CA) and cells were transfected with
50ul Lipofectamine 2000 (Invitrogen) and 20 µg plasmid DNA per plate. Lipid-
DNA complexes were incubated 4 to 5 hours on cell monolayers at 37ºC/5% CO2 after which transfection medium was replaced with complete medium. Cells were then cultured for an additional 24-36 h before experiments.
Isolation of crude membranes. Mouse tissue or cultured cells were isolated, washed with 25 mM imidazole, 1 mM EDTA, 250 mM sucrose and protease inhibitor cocktail (Sigma) and manually homogenized with 25 strokes of a prechilled Dounce homogenizer. Nuclei, unbroken cells and mitochondria were separated by centrifuging at 6000 x g for 5 minutes. The supernatant was collected and saved, while the pellet was homogenized again and centrifuged.
The two supernatants were combined and centrifuged at 125,000 x g to pellet crude microsomal membranes. The pellets were resuspended in 25 mM imidazole, 1 mM EDTA, 20 mM NaCl and stored at 4ºC.
Immunoblot analysis of FXYD5 and SGK1. Crude membrane (10-20 µg) were dissolved in 30µl 2x Laemmli buffer, incubated at 37ºC for 20 minutes, loaded
164 onto 12% SDS-PAGE and resolved at 100 V for 1.5 hr. Proteins were transferred via semi-dry blotting onto nitrocellulose (Biorad), blocked with 1% BSA and incubated with either rabbit polyclonal anti-FXYD5 562 (1:250) antibody, mouse anti-M2 Flag tag (1 µg/ml, Sigma) or mouse anti-SGK1 (1 µg/ml, Upstate
Biotech) in 1% BSA/Tris-buffered saline for 1 hr at room temperature. The blot was then incubated with species specific secondary antibodies (1:4000) conjugated to HRP and visualized using Pierce’s Supersignal Pico West detection kit.
Indirect immunofluorescence. MDCK cells were cultured in 4-well chamber slides
(Permanox), washed twice with phosphate-buffered saline (PBS) and fixed for 5 minutes in 100% ice-cold methanol. Slides were then rinsed twice with PBS, blocked for non-specific antibody binding by incubating the cells in 4% bovine serum albumin for 1 hour (Invitrogen, Carlsbad, CA). Primary rabbit anti-ECS
(Flag) antibody (2 µg/ml) (Bethyl Labs, Montgomery, TX) was diluted in 1%
BSA/PBS, incubated on monolayers for 45 minutes and aspirated. Monolayers were washed twice with PBS and then incubated in 1% BSA/PBS containing goat anti-rabbit IgG Alexa-Fluor 568 (1:250, Invitrogen, Carlsbad, CA) for 45 minutes.
Cells were then washed twice, the nuclei counterstained with Hoechst 33342 dye
(Invitrogen, Carlsbad, CA) and mounted with Fluormount-G. Slides were allowed to dry overnight and immunofluorescent localization was assessed on a Zeiss
200M Axiovert inverted microscope, with a DG4 switchable fluorescent light source (Sutter Instrument Company, Novato, CA) and a 12-bit CoolSnap HQ
165 camera (Roper Scientific, Tucson, AZ) under control of MetaMorph v 6.2
(Molecular Devices, Sunnyvale, CA. Images were obtained with a 63X numerical aperture 1.3 fluar lens using excitation and emission filter passbands of 260 ±
20nm and 645 ±30 nm, respectively. Typical exposure times for individual frames were 150 ms.
Immunoprecipitation of FXYD5-Flag. Immunoprecipitation was performed as previously described [119]. Briefly, crude membrane protein (125 µg) from HEK
293 cells was resuspended in 10 mM RbCl and 200 µM ouabain. Aliquots of solubilized membrane were incubated with 1 µg M2 Flag antibody or non-specific
IgG at 4ºC for 3 hours. 50 µl of Protein G agarose beads were added and incubated overnight on a rocking platform at 4ºC. The beads were washed two times with 25 mM imidazole, 1mM EDTA, 100 mM RbCl, 200 µM and 0.2mg/ml
C12E10 and either washed twice again in assay dilution buffer 2 (ADB2) for kinase assay preparation or resuspended in 50 µl Laemmli buffer, incubated at 37ºC for
20 minutes and loaded onto 12% SDS-PAGE. After semi-dry transfer to nitrocellulose, proteins were blocked with 1% BSA and incubated with antibodies raised against the M2 Flag tag (1mg/ml) (Sigma, St. Louis, MO) in 1% BSA/Tris- buffered saline for 1 hr at room temperature. The blot was then incubated with species specific secondary antibodies (1:4000) conjugated to HRP and visualized using Pierce’s Supersignal Pico West detection kit.
166 Kinase assay. Peptides corresponding to residues (160-178) of mouse and human FXYD5 were synthesized by the Cleveland Clinic Lerner Research
Institute on an Applied Biosystems Model 431A synthesizer. To assess the ability of various kinases to phosphorylate the two serine residues in the carboxy terminus of FXYD5, Ser163 and Ser170 were separately mutated to alanine.
Peptides were reconstituted in water and protein concentration was determined using the BioRad Protein Assay (BioRad). Kinase phosphorylation was assessed using the SGK1(Δ1-59,S422D) and PKC Kinase Assay kits (Upstate). Briefly, mouse or human FXYD5 peptide was diluted in Assay Dilution buffer II (ADB2) to a final concentration of 500 µM, mixed with 10 µCi [γ-32P]ATP and incubated with either 50 ng SGK1 for 10 minutes at 30ºC. 25µl aliquots were then spotted on
P81 phosphocellulose paper, washed three times with 0.75% phosphoric acid, washed once with acetone and counted in a scintillation counter to compare [γ-
32P] incorporation. Assays were performed in triplicate and repeated. To assess the ability of SGK1 to phosphorylate mature FXYD5 protein, HEK293 cells were transfected with pKCERhF5kFlag, pKCERhF5kFlagS163A or pKCERhF5kFlagS170A and utilized for immunoprecipitation analysis as described above. 20 µl HEK293 FXYD5-Flag immunoprecipitate was with ADB2 and phosphorylated by SGK1 as described above.
86Rb+ uptake. Ouabain-inhibitable uptake of 86Rb+ was used as a measure of K+ transport by Na,K-ATPase in HEK 293 cells. Monolayers were grown to 80% confluency in 12 well tissue culture plates (Costar) and transfected as described above. Transfection efficiency was assessed by GFP fluorescence in a cell
167 monolayer transfected in parallel. On day 2, monolayers washed once and incubated for 15 minutes in (mM): 140 NaCl, 1 MgCl2, 1 CaCl2, 1 RbCl, 2
Na2PO4, 5 glucose, 5 HEPES and 200 µM bumetanide (pH 7.4) and in the presence or absence of 2 mM ouabain. Uptake was initiated with the addition of
0.75 µCi 86Rb+ and monolayers were incubated for 15 minutes in a 37ºC water bath. All solutions were warmed to 37ºC prior to use. Initial experiments determined that uptake was linear for 30 minutes (data not shown). Uptake was terminated by aspiration of the incubation buffer and subsequent washes with (in mM) 140 NaCl, 1 MgCl2, 1 CaCl2, 4 RbCl, 2 BaCl2, 5 HEPES, 5 glucose, 200 µM bumetanide. The cells were allowed to air dry, solubilized in 0.1mM NaOH and
0.5% SDS and the amount of 86Rb+ associated with the monolayers was assessed by liquid scintillation spectrometry. Na,K-ATPase associated 86Rb+ uptake was calculated as the difference between total and ouabain-sensitive
86Rb+ uptake.
Statistics. Microsoft Excel was used for calculating Student’s T-test. Sigma Stat was used to calculate ANOVA statistics using the Student-Neuman-Kuels regression analysis for pairwise comparisons. P<0.05 was used to declare statistical significance unless otherwise noted.
168
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