Regulation of renin gene expression by CTCF, Nr2f2, Nr2f6, Nr4a1

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Spring 2011

Regulation of renin gene expression by CTCF,Nr2f2, Nr2f6, Nr4a1 and maintenance of the reninexpressing cellEric Thomas WeatherfordUniversity of Iowa

Copyright 2011 Eric Thomas Weatherford

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Recommended CitationWeatherford, Eric Thomas. "Regulation of renin gene expression by CTCF, Nr2f2, Nr2f6, Nr4a1 and maintenance of the reninexpressing cell." PhD (Doctor of Philosophy) thesis, University of Iowa, 2011.https://doi.org/10.17077/etd.f8f6i6xy

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REGULATION OF RENIN GENE EXPRESSION BY CTCF, NR2F2, NR2F6, NR4A1

AND MAINTENANCE OF THE RENIN EXPRESSING CELL

by

Eric Thomas Weatherford

An Abstract

Of a thesis submitted in partial fulfillment of the requirements for the Doctor of

Philosophy degree in Molecular Physiology and Biophysics in the Graduate College of

The University of Iowa

May 2011

Thesis Supervisor: Professor Curt D. Sigmund

1

ABSTRACT

The renin angiotensin system (RAS) is critical for the regulation of blood

pressure, electrolyte/fluid, and metabolic homeostasis. Regulation of the RAS is

important in the development and treatment of hypertension. As part of the rate-limiting

step in a cascade of events ending in the production of angiotensin II, renin is a major

regulator of the RAS. Its expression is localized to the juxtaglomerular (JG) cells of the

JG apparatus where it is exquisitely located to respond to various physiological cues.

Understanding the regulation of renin expression and development of the juxtaglomerular

cells is critical. Two regulatory elements, the enhancer and proximal promoter, have

been found to be important in controlling cell- and tissue- specific baseline expression of

the renin gene. Within the enhancer is a hormone response element (HRE) which confers

a high level of activity to the enhancer. Nuclear receptors that bind this element have

been found to bind the HRE and regulate renin promoter transcriptional activity. We

have previously characterized the role of the orphan nuclear receptor Nr2f6 as a negative

regulator of renin expression that mediates its effects through the HRE. However, gel

shift assays indicate that there are other transcription factors binding this element. We

have identified other orphan nuclear receptors that regulate renin expression. The first,

Nr2f2 acts as a negative regulator of renin promoter activity but does not appear to affect

baseline expression of the endogenous renin gene. The other, Nr4a1, is a positive

regulator of renin expression, but it does not appear to mediate its effects through the

HRE.

The transcriptional regulation of gene expression is controlled by regulatory

elements separated by large distances from promoters. We and others have found that

short transgenes of the human renin (hREN) locus are not sufficient to protect them from

positional effects that can be exerted upon them by neighboring regulatory elements. We

discovered a random truncation in a large genomic construct of the hREN gene that

2

resulted in ubiquitous expression of renin not seen with the intact form. By locating the

genomic insertion site of that transgene in the Zbtb20 gene, we found that the hREN

promoter had come under control of that gene’s regulatory elements. The gene

downstream of renin however maintained its tissue-specific expression. We found that

CCCTC-binding factor (CTCF) bound to chromatin in and around the renin locus. The

presence of CTCF suggests that insulator elements are present in the renin locus, and

their loss likely explains the results above.

Finally, we assessed the role of microRNAs in the development of renin

expressing cells in the mouse kidneys by cell-specific deletion of the processing enzyme

Dicer. This resulted in reduction of renin expression and a decrease in the number of

renin expressing cells in the kidney. Mice were hypotensive and had several kidney

abnormalities including a hypertrophied vasculature and striped fibrosis. These results

indicate that Dicer and the miRNAs it processes are critical for the development and

maintenance of renin expressing cells that contribute to normal kidney development.

Abstract Approved: ____________________________________ Thesis Supervisor

____________________________________ Title and Department

____________________________________ Date

REGULATION OF RENIN GENE EXPRESSION BY CTCF, NR2F2, NR2F6, NR4A1

AND MAINTENANCE OF THE RENIN EXPRESSING CELL

by


A thesis submitted in partial fulfillment of the requirements for the Doctor of

Philosophy degree in Molecular Physiology and Biophysics in the Graduate College of

The University of Iowa

May 2011

Thesis Supervisor: Professor Curt D. Sigmund

Graduate College The University of Iowa

Iowa City, Iowa

CERTIFICATE OF APPROVAL

_______________________

PH.D. THESIS

_______________

This is to certify that the Ph.D. thesis of


has been approved by the Examining Committee for the thesis requirement for the Doctor of Philosophy degree in Molecular Physiology and Biophysics at the May 2011 graduation.

Thesis Committee: ___________________________________ Curt D. Sigmund, Thesis Supervisor

___________________________________ Michael G. Anderson

___________________________________ Michael D. Henry

___________________________________ Scott Moye-Rowley

___________________________________ Kamal Rahmouni

___________________________________ Thomas J. Schmidt

To my members of the “Greatest Generation”

ii

ABSTRACT

The renin angiotensin system (RAS) is critical for the regulation of blood

pressure, electrolyte/fluid, and metabolic homeostasis. Regulation of the RAS is

important in the development and treatment of hypertension. As part of the rate-limiting

step in a cascade of events ending in the production of angiotensin II, renin is a major

regulator of the RAS. Its expression is localized to the juxtaglomerular (JG) cells of the

JG apparatus where it is exquisitely located to respond to various physiological cues.

Understanding the regulation of renin expression and development of the juxtaglomerular

cells is critical. Two regulatory elements, the enhancer and proximal promoter, have

been found to be important in controlling cell- and tissue- specific baseline expression of

the renin gene. Within the enhancer is a hormone response element (HRE) which confers

a high level of activity to the enhancer. Nuclear receptors that bind this element have

been found to bind the HRE and regulate renin promoter transcriptional activity. We

have previously characterized the role of the orphan nuclear receptor Nr2f6 as a negative

regulator of renin expression that mediates its effects through the HRE. However, gel

shift assays indicate that there are other transcription factors binding this element. We

have identified other orphan nuclear receptors that regulate renin expression. The first,

Nr2f2 acts as a negative regulator of renin promoter activity but does not appear to affect

baseline expression of the endogenous renin gene. The other, Nr4a1, is a positive

regulator of renin expression, but it does not appear to mediate its effects through the

HRE.

The transcriptional regulation of gene expression is controlled by regulatory

elements separated by large distances from promoters. We and others have found that

short transgenes of the human renin (hREN) locus are not sufficient to protect them from

positional effects that can be exerted upon them by neighboring regulatory elements. We

discovered a random truncation in a large genomic construct of the hREN gene that

iii

resulted in ubiquitous expression of renin not seen with the intact form. By locating the

genomic insertion site of that transgene in the Zbtb20 gene, we found that the hREN

promoter had come under control of that gene’s regulatory elements. The gene

downstream of renin however maintained its tissue-specific expression. We found that

CCCTC-binding factor (CTCF) bound to chromatin in and around the renin locus. The

presence of CTCF suggests that insulator elements are present in the renin locus, and

their loss likely explains the results above.

Finally, we assessed the role of microRNAs in the development of renin

expressing cells in the mouse kidneys by cell-specific deletion of the processing enzyme

Dicer. This resulted in reduction of renin expression and a decrease in the number of

renin expressing cells in the kidney. Mice were hypotensive and had several kidney

abnormalities including a hypertrophied vasculature and striped fibrosis. These results

indicate that Dicer and the miRNAs it processes are critical for the development and

maintenance of renin expressing cells that contribute to normal kidney development.

iv

TABLE OF CONTENTS

LIST OF TABLES............................................................................................................ vii

LIST OF FIGURES ......................................................................................................... viii

CHAPTER 1 GENERAL INTRODUCTION .....................................................................1 History and Background of the Renin-Angiotensin System.............................2 Regulation of Renin Expression .......................................................................6

Transcriptional Regulation of Renin .........................................................6 Post-transcriptional regulation.................................................................11

Regulation of Renin Secretion........................................................................13 Development of the Renin Expressing Cell....................................................15

CHAPTER 2 CONTROL OF RENIN EXPRESSION BY NUCLEAR RECEPTORS.............................................................................................20

Introduction.....................................................................................................20 Methods ..........................................................................................................22

RT-PCR ...................................................................................................22 cDNA Expression Plasmids ....................................................................22 Luciferase Assay .....................................................................................22 Orphan Nuclear Receptor Knockdown ...................................................23 Electrophoretic Mobility Shift Assay and Supershift Assay...................24 DNA Affinity Purification Assay............................................................24 Chromatin Immunoprecipitation .............................................................25 Immunofluorescence ...............................................................................25

Results.............................................................................................................26 Discussion.......................................................................................................31

CHAPTER 3 CONTROL OF THE RENIN LOCUS BY CTCF.......................................46 Introduction.....................................................................................................46 Methods ..........................................................................................................47

Generation of Kidney Enhancer-deficient PAC160 Transgenic Mice.........................................................................................................47 Transgene insertion mapping ..................................................................48 RNA Isolation and RT-PCR....................................................................48 In vitro Transcription Translation ...........................................................49 EMSA......................................................................................................49 Chromatin Immunoprecipitation .............................................................50

Results.............................................................................................................50 Discussion.......................................................................................................54

CHAPTER 4 DICER MAINTAINS THE RENIN CELL PHENOTYPE.........................66 Introduction.....................................................................................................66 Methods ..........................................................................................................67

Generation of Dicer KO mice..................................................................67 Histological analysis and immunostaining..............................................68 RNA extraction and quantitative RT-PCR (qRT-PCR) analysis ..............68

v

Tail cuff and Telemetry BP measurements .............................................68 Results.............................................................................................................69 Discussion.......................................................................................................70

CHAPTER 5 GENERAL DISCUSSION ..........................................................................76 Renin Expression and Maintenance of the Renin Cell Phenotype .................76

Summary of Results.................................................................................76 Future Directions .....................................................................................77

REFERENCES ..................................................................................................................81

vi

LIST OF TABLES

Table 2-1. Primers and probes for gene expression analysis. ...........................................37

Table 2-2. Summary of data from RT-PCR and microarray analysis of As4.1 mRNA..............................................................................................................39

Table 3-1. Primers and probes for determining gene expression, transgene insertion mapping, EMSA, and ChIP analysis................................................................58

vii

LIST OF FIGURES

Figure 1-1. Alignment of mouse and human enhancer.....................................................19

Figure 2-1. Expression of orphan nuclear receptors in As4.1 cells. .................................38

Figure 2-2. Immunofluorescence for Nr2f2, Nr2f6, and Nr4a1. ......................................40

Figure 2-3. Luciferase activity of 4.1kP-luc when Nr2f2, Nr2f6, or Nr4a1 are overexpressed or knocked down....................................................................41

Figure 2-4. EMSA analysis of in vitro translated Nr2f2, Nr2f6, and Nr4a1. ...................42

Figure 2-5. Characterization of Nr2f2, Nr2f6, and As4.1 nuclear proteins to the HRE................................................................................................................43

Figure 2-6. Renin, Nr2f2, Nr2f6, and Nr4a1 expression following orphan receptor knock down in As4.1 cells. ............................................................................44

Figure 2-7. Expression of other potential target genes following orphan receptor knock down....................................................................................................45

Figure 3-1. Stucture of the PAC160 and ∆KE6 transgenes and renin expression............59

Figure 3-2. Mapping the ∆KE6 insertion site in the mouse genome. ...............................60

Figure 3-3. Detection of fusion transcripts in ∆KE6 mice................................................61

Figure 3-4. Genome structure and transcripts of ∆KE6 mice...........................................62

Figure 3-5. RNase protection assay identifying fusion versus hREN transcripts.............63

Figure 3-6. EMSA analysis of identified CTCF binding sites..........................................64

Figure 3-7. ChIP analysis for the binding of chromatin around the ∆KE6 transgene. .....65

Figure 4-1. Expression of Ren1 and Ren2 in mouse kidneys of control and knockout mice................................................................................................73

Figure 4-2. Blood pressure measurements in control and Dicer knockout mice. .............74

Figure 4-3. Histology of control and Dicer knockout mice. .............................................75

viii

1

CHAPTER 1

GENERAL INTRODUCTION

The renin-angiotensin system (RAS) or renin-angiotensin-aldosterone system

(RAAS) maintains homeostasis by regulating blood pressure and fluid-electrolyte

balance. How the system accomplishes the modulation of these endpoints is as diverse as

the tissues responding to angiotensin II (Ang II). Ang II can regulate blood pressure by

directly stimulating vasoconstriction in resistance vessels. It also has direct and indirect

actions in the kidney, and its regulation of fluid-electrolyte homeostasis can indirectly

influence blood pressure by regulating extracellular volume. Angiotensin II stimulates

aldosterone release from the adrenal glands leading to Na+ reabsorption in the kidney, a

driving force for H2O retention. Ang II can also directly stimulate sodium reabsorption

by binding to Ang II receptors in the tubular segments of the late nephron. Furthermore,

Ang II can stimulate the thirst response, salt appetite, and vasopressin release by actions

in the brain. Sympathetic outflow is also stimulated by Ang II and that can affect other

levels of blood pressure regulation like chronotropic and ionotropic events in the heart.

Furthermore, β-adrenergic stimulation through the sympathetic nervous system is one

way to activate the RAS by stimulating renin secretion from the kidney. Initially

believed to be a purely paracrine system, it is widely accepted that it functions in an

autocrine or “intracrine” manner in several tissues. Initially simplistic and straight

forward, the complexity of the RAS has expanded significantly since discovery of the

founding molecule renin.

Renin serves as the foundation of this system because it is involved in the rate-

limiting step of a cascade of events leading to physiological changes. It is primarily

produced, stored, and secreted from the juxtaglomerular (JG) cells of the kidney. These

cells make up approximately 0.01% of the total cell population of the kidney, yet account

for the bulk of renin produced body-wide. They are exquisitely positioned adjacent to

2

both the vascular and tubule components of the kidney to respond to blood pressure and

electrolyte changes of which the RAS is responsible for regulating.

History and Background of the Renin-Angiotensin System

Study of the RAS was initiated by the discovery of renin by Robert Tigerstedt and

Per Bergman over 100 years ago. They discovered that kidney extracts were able to

induce increases in blood pressure when injected into rabbits. They hypothesized that the

kidney produced a vasoactive substance and were able to localize its production to the

kidney cortex. Because of the source they called the substance renin. Their discovery

was largely ignored until the work of Henry Goldblatt published in 1934. He noticed that

patients who died of hypertension often had a narrowing of renal blood vessels.

Goldblatt was able to show that clipping of renal arteries resulted in hypertension in

experimental animals (1). He too concluded that the ischemic kidney must produce a

substance with vasoactive properties (2). This spurred new interest in the work of

Tigerstedt and whether renin was responsible for the hypertension in Goldblatt’s

experiments.

The mechanism of increased blood pressure produced by Goldblatt’s ischemic

kidney was pursued by two separate groups simultaneously. One was led by Irvine Page

in the United States and the other by Eduardo Braun-Menendez in Argentina. Page’s

group noted that there was a diminished response to renin in isolated perfused dog tail

preparations unless plasma was included with the perfusate (3). This led to the conclusion

that renin was not the vasoactive substance, but acted on another plasma peptide.

Menendez’s group demonstrated that renal vein blood from ischemic kidneys produce a

strong pressor response in nephrectomized dogs. Further work resulted in the

characterization of another vasoactive molecule that had very acute pressor action unlike

the prolonged effect of renin (4). One group named this substance “hypertensin”, the

other “angiotonin”, which were combined to form the name as we know it today,

3

“angiotensin”. Furthermore, renin was recognized as a proteolytic enzyme released from

the kidney that acted on a plasma substrate, hypertensinogen (angiotensinogen). In 1942,

Leloir was able to show that this substrate is produced in the liver, and in 1954, Leonard

Skeggs was able to show that angiotensin existed in two forms, angiotensin I (Ang I) and

II (Ang II) (5-7). Skeggs was subsequently able to show that Ang I converted by

angiotensin converting enzyme (ACE) to Ang II was responsible for the increases in

blood pressure (8). About four years later it was determined that Ang II was able to

stimulate aldosterone release (9). The last piece to the puzzle came in 1970 when

angiotensin receptors were identified in various tissues (10). That completed the pathway

as we know it today- renin release from the kidney cleaves angiotensinogen to Ang I

which is further processed by ACE producing the active peptide Ang II that can stimulate

vasoconstriction and aldosterone release by binding to angiotensin receptors.

The RAS soon became a system that was considered to be attractive as a drug

target for controlling high blood pressure (hypertension). The first drug utilized to target

the RAS was the ACE inhibitor captopril in 1977. This drug was based on a peptide

found in the Brazilian pit snake, Bothrops jararaca. The existence of such a peptide was

first described by Sergio Ferreira in 1965 (11). In 1968 Y.S. Bakhle showed that ACE

from the dog lung was inhibited by peptides from this snake’s venom (12). David

Cushman, a biochemist, and Miguel Ondetti, a chemist, collaborated to develop captopril

based on structural combinations of the peptide from pit snake venom and L-

benzylsuccinic acid (13). Inhibitors of ACE are now some of the most prescribed anti-

hypertensive drugs prescribed today. Today, other ACE inhibitors have been developed

as well as Ang II receptor antagonists and renin inhibitors. Not only was captopril the

first drug to target the RAS, but it renewed interest in its role in controlling blood

pressure.

The RAS has expanded greatly in terms of the number of angiotensin peptides

generated, receptors involved, and localization of its components. There has been the

4

discovery of a second ACE (ACE2) that is important in the steps that generate Ang-(1-7),

another peptide hormone of the RAS (14, 15). The binding of Ang 1-7 to its receptor (the

Mas receptor) appears to initiate cell responses that antagonize those of Ang II (16).

Renin is secreted in both an inactive (prorenin) and active form. Recently, a

prorenin/renin receptor (PRR) has been identified and characterized to bind prorenin.

When prorenin binds to the receptor, its active site is unmasked allowing it to cleave

angiotensinogen and generate Ang II (17). A signal transduction cascade that results in

ERK1/2 activation is also initiated upon prorenin/renin binding. Thus, a new hypothesis

is that renin is not only an enzyme responsible for the ultimate production of Ang II, but

is also a ligand activating intracellular signaling by binding its receptor. The PRR may

play an important role in localized tissue Ang II production and local RASs.

It is now well accepted that several organs express components of the RAS. Of

particular importance is their expression in key cardiovascular tissues such as the kidney,

heart, and brain. It has become increasingly apparent that these local tissue RASs are

involved in cardiovascular disease progression. The potential importance of these tissue

RASs is highlighted by the fact that ACE inhibitors and ARBs can lower blood pressure

even in patients with low or normal plasma renin levels, which points to inhibition at the

level of local systems (18, 19).

The primary sites of renin synthesis and release are the juxtaglomerular (JG) cells

of the JG apparatus in the kidney. However, other bonafide sites of renin expression

exist, with the brain and lung expressing specific novel transcriptional isoforms of the

renin mRNA containing exon-1b and exon-1c, respectively, in place of exon-1a (20).

These isoforms are probably regulated by different promoters since the expression of

exon-1b is retained despite deletion of the 500 bp of DNA surrounding exon-1a that

includes the classic renin promoter (21). These two isoforms are most likely intracellular

forms of renin and probably participate exclusively in localized tissue RAS signaling.

Overexpression of the RAS specifically in the brain of mice leads to hypertension and

5

increases in metabolic rate with suppression of the peripheral RAS (22-24). When renin

is overexpressed in the proximal tubule of the kidney it leads to hypertension without

stimulation of the peripheral RAS (25). These two animal models are just two examples

that underscore the significance of tissue RASs in hypertension.

Hypertension is a major risk factor for cardiovascular disease and lowering blood

pressure in patients has proven to significantly reduce risk for developing cardiovascular

morbidity and mortality (26-28). In human patients, hypertension is defined as a systolic

blood pressure greater than 130 mmHg and/or a diastolic greater than 90 mmHg. A

major controller of blood pressure homeostasis is the RAS. Renin catalyzes the

conversion of angiotensinogen to angiotensin I in the rate-limiting step of the RAS, and

angiotensin I is ultimately cleaved by angiotensin converting enzyme (ACE) to produce

angiotensin II. This molecule is the primary blood pressure regulator of the RAS. It

stimulates vasoconstriction, sodium reabsorption, salt appetite and thirst, vasopressin

release, and the sympathetic nervous system. The importance of the RAS in controlling

blood pressure, hypertension, and development of cardiovascular disease is revealed by

the effectiveness of ACE inhibitors and angiotensin receptor blockers (ARBs) as

antihypertensive agents (29, 30). In fact, the renin inhibitor Aliskiren was recently

approved for clinical use. The RAS also plays an important role in Mendelian forms of

hypertension, and renin gene polymorphisms have been associated with hypertension

(31-33). In conjunction with control of blood pressure homeostasis, the RAS, and

particularly renin, plays a significant role in kidney development and is critical

postnatally (34-37). Specifically, secreted renin is critical for life as its genetic ablation

leads to postnatal lethality (21). Because renin is involved in the most regulated and rate-

limiting step of the RAS, understanding its regulation is critical to resolving the

complexity of hypertension and cardiovascular disease.

6

Regulation of Renin Expression

Transcriptional Regulation of Renin

The mouse and human renin genes are found on chromosome 1 and share about

78% homology. Certain strains of mice (e.g. C57BL/6 & BALB/c) carry one renin gene

whereas others (e.g. DBA/2J & 129SvJ) have two as a result of gene duplication. Renin

in one gene strains is designated Ren-1c, and in two gene strains, renin loci are termed

Ren-1d and Ren-2. The coding regions of Ren-1c and Ren-1d are 99% homologous while

Ren-1c&d are 97% homologous to Ren-2. One gene strains more faithfully recapitulate

the genetic state in humans. Therefore, the majority of evidence presented here will

focus on the Ren-1c gene.

Initial studies into the transcriptional regulation of renin involved the use of non-

renin expressing cells and transgenic mice. Data from non-renin expressing cells are

probably of limited use because the renin gene was not studied in its usual environment

of trans-acting factors. Transgenic mice, however, provided evidence that ~4.6 kb of

Ren-2 upstream sequence was sufficient to drive tissue- and cell-specific renin expression

of the T-antigen coding sequence (38). Now, through the use of endogenous renin

expressing juxtaglomerular derived mouse As4.1 and human lung carcinoma Calu-6

cells, we have a better understanding of the trans- and cis-factors involved in the

regulation of the renin gene (39, 40).

Evidence from As4.1 cells has identified two elements within the Ren-1c gene that

are critical for its control. A proximal promoter element (PPE, -197 to -50 bp) and

enhancer (-2866 to -2625 bp) work in conjunction to direct renin expression. Inclusion of

the enhancer in reporter constructs results in a ~50-fold increase in promoter activity in

an orientation- and position-independent manner (41). The mREN enhancer shares 71%

homology with a human renin (hREN) enhancer that lies ~11 kb upstream of the start site

(Figure 1-1). Within the enhancer there are three identified binding sites that are 100%

7

identical between mouse and human. Mutation of any one of these sites results in a

dramatic decrease in enhancer activity (42, 43). Element d (Ed) is a cAMP responsive

element (CRE) which has been found not only to bind cAMP responsive element binding

protein (CREB) and cAMP responsive element modulator (CREM), but also nuclear

factor kappa B (NfκB) (44). A second site, termed element e (Ee), is an E-box found to

bind upstream stimulatory factors 1 and 2 (USF-1/2) (42). The last binding site,

element c (Ec), is a hormone response element (HRE) (45). This site is of particular

interest because it is the half-site of a TGACCT repeat, that in combination with element

b (Eb) of the mouse enhancer makes up a larger HRE. Additionally, the HRE can bind

EAR2 and vitamin D receptor (VDR) that have a negative effect on the enhancer (43,

46). However, the effects of VDR appear to be controlled through DNA independent

transrepression of CREB binding to the CRE (47). Within the second half-site of the

HRE, the human enhancer contains a A>G single nucleotide polymorphism (SNP) that

reduces enhancer activity significantly and may contribute to the dramatic differences

seen in renin levels between mice and humans (~1000 vs. 3 ng Ang I/ml/hr) (48).

The remaining portion of the enhancer contains six binding sites that are less

conserved yet may still be important for enhancer function. Of those six sites, four are

NFI binding sites and one is a SP1/3 site (49). When the binding sites are mutated

individually they have generally modest effects on promoter activity. When all the sites

are mutated there is approximately a 90% reduction. However, their action appears to be

dependent on the presence of the proximal portion of the enhancer discussed above (42).

In addition to the enhancer and its transcription factors, the PPE is a key element

of the renin gene. Within the PPE, there is a binding element named the renin proximal

promoter element-2 (RP-2) that binds the HOXB9/D10-PBX1b-MEIS/PREP1 complex

and is involved in baseline and cAMP induced renin expression (42, 50). When the RP-2

element is deleted in transgenic mice, renin expression in the kidney is lost, but not in

other tissues (51, 52). This element is thus critical for tissue-specific expression of renin.

8

The HOXB9/D10-PBX1b complex appears to mediate cAMP responsiveness through

binding to CREB since it cannot bind to the site by itself. Furthermore, this complex can

cooperate with Ets and Notch1/CBF1 to activate rat renin promoter activity (53). That

HoxD10 and Notch1 direct promoter activity and are well known developmental

regulators, suggests renin may help determine cell fate. In fact, retinoblastoma

susceptibility protein (RB) can stimulate renin expression in human embryonic kidney

cells through the same binding site as HOXB9/B10 (54). This is of note since human

embryonic kidney cells do not normally express renin. While the HOX and PBX binding

sites appear to be critical for promoter activity across species, other possible cis-elements

have been identified via DNaseI footprinting assays of the human promoter (55, 56). One

of these sites is a CRE that binds ATF1-CREB heterodimer, but it is poorly conserved

across species (57). The other site is for the orphan nuclear receptor Nr2f2 (Coup-tfii,

ARP-1) that negatively regulates human promoter activity (58). However, this site is not

well conserved across species and it has not been determined whether the corresponding

site in the mouse or rat promoters is still capable of binding Nr2f2. The promoter is no

doubt critical for controlling renin, but some evidence reveals that it may only direct cell-

specific expression by interacting with the appropriate combination of cis-elements (59-

61). Initial data suggested that the enhancer identified in As4.1 cells might be the

element to fill this role, but as will be discussed subsequently, this is probably not the

only element responsible.

Renin is under tight transcriptional and post-transcriptional control. Physiological

regulators of renin include perfusion pressure, β-adrenergic stimulation, angiotensin II

negative feedback, and tubuloglomerular feedback. The major intracellular pathways

activated/inhibited by these regulators involve the cAMP pathway and Ca2+. It now

appears that these two second messengers are connected in their modulation of renin by

Ca2+-inhibitable adenylate cyclases V and VI (62, 63). Of note is the observation that

Ang II can inhibit renin exocytosis in isolated perfused kidneys when stimulated through

9

adenylate cyclase with isoproterenol, but not stimulation by 8Br-cAMP. This same study

revealed that angiotensin II activates increases in intracellular Ca2+ in cultured JG cells.

This study provides a direct link to a physiological negative regulator and the signaling

mechanism involved. It does not test effects at the transcript level, but increases in

intracellular Ca2+ have previously been shown to inhibit renin via transcriptional and

post-transcriptional mechanisms (64). Cyclic AMP response element binding protein

(CREB) is a transcription factor target of this pathway and CRE binding sites in the renin

promoter and enhancer play an integral role in renin transcription. However, there exist

CREB independent pathways that activate human promoter activity through the CRE (42,

50, 57). Beyond the cAMP pathway, there is less known about the pathways that link

physiological cues to increases or decreases in renin expression. Evidence does suggest

an important role for the enhancer in mediating negative regulation in response to

cytokines and vitamin D, while regulating positive stimulation by retinoic acid (45, 46,

65). Taken together, the evidence provided above would suggest that the enhancer is a

master regulator of renin.

Studies in vivo where the enhancer has been specifically deleted points us in a

new direction. Two separate models have been developed to study the effects of deleting

the renin enhancer (66, 67). These studies imply that both the human and mouse

enhancers are important for baseline expression, but not for cell- and tissue-specific

expression. The renin enhancer knockout (REKO) mouse provided evidence that the

enhancer is critical for renin expression and regulation, but it fails to use more sensitive

methods for renin detection and specificity of expression. In a separate study using the

REKO mouse, Markus et al. show that there are modest decreases in kidney tissue renin

activity and mRNA (68). Plasma renin activity (PRA) and concentration (PRC) were

also reduced, but did not reach statistical significance. The lack of a more significant

response may be due to positive feedback on renin exerted by the blood pressure decrease

observed in these mice. This region may be of critical importance in the salivary gland

10

and adrenals, as its deletion resulted in almost complete loss of renin in these tissues.

Despite a modest effect on baseline renin, when challenged with a low salt diet combined

with enalapril administration, REKO mice had significantly blunted increases. Perhaps

most interesting was an almost complete absence of kidney renin protein. Although the

mRNA levels in the kidney still increased, it was insufficient to replenish kidney renin

stores in response to the physiological change. This study did not address whether a

partial response was still sufficient to maintain the baseline blood pressure. These results

support the role of the enhancer in regulating not only the baseline activity of the

promoter, but also the transcriptional response to physiological cues and stressors.

Our lab focused on the human enhancer by precisely deleting the enhancer within

a PAC transgene carrying the entire renin gene plus 75 kb upstream and 70 kb

downstream sequences (69). This model is advantageous because of the species-

specificity of the renin-angiotensinogen reaction. Unlike the REKO mouse, knock out of

the human renin enhancer in the PAC model and subsequent decrease in renin expression

would have no impact on the blood pressure because human renin cannot cleave mouse

angiotensinogen. This allows for isolating the effects of deleting the enhancer without

changing the normal physiology of the mouse. Our data showed that normal renin

expression in transgenic mice was retained, albeit at lower levels, and responded

appropriately to captopril treatment (ACE inhibition) as well as angiotensin II infusion.

Our data indicate that the enhancer is required to regulate baseline renin expression, but

is dispensable for cell-specific expression or response to physiological cues.

Data from Ken Gross’s lab using a BAC transgenic with the renin coding region

replaced with GFP largely recapitulates our results (52). Their model targets the 3’

portion of the enhancer containing the three binding sites discussed earlier that are most

critical for its activity (Ee, Ed, Ec). Deletion of this region along with Eb and Ea led to a

large reduction in baseline GFP expression. Unlike our results, they observed a

diminished increase in GFP when mice were given captopril. This may represent a

11

mouse promoter specific response since this is in agreement with the REKO mouse.

Importantly, their data provide in vivo evidence supporting the luciferase experiments

that localize the bulk of enhancer function to the 3’ region containing elements Ea

through Ee.

There is another renin enhancer in the human gene at position -5777 to -5552 that

we have termed the chorionic enhancer (CE). This enhancer was identified by transient

transfection of reporter constructs in choriodecidual cells (70). It induced a 59-fold

increase in promoter activity in choriodecidual cells, but a much less robust 6-fold

increase in As4.1 and Calu-6 cells. There is a SNP downstream of this enhancer that

mediates differential activation of the promoter in transient transfection reporter gene

analysis in choriodecidual cells (71). The region containing the SNP does bind proteins

from choriodecidual nuclear extracts, but the SNP has little impact in patients (72). What

transcription factors and binding sites regulate promoter activity through this element

remains to be determined. Our lab deleted this element in PAC transgenic mice and

found no change in the regulation of kidney or placental renin expression (73).

Additionally, its deletion had no effect on the expression of the nearby KISS gene. This

was tested because the KISS gene is expressed highly in placenta, and thus we

hypothesized that perhaps the CE is not an upstream enhancer for renin but a downstream

enhancer of KISS. These data would suggest that this element plays no role in renin

expression.

Post-transcriptional regulation

The first analysis of post-transcriptional regulation of renin was performed in

native juxtaglomerular cells. Renin mRNA half-life was measured to be 3 hours, but

when treated with forskolin it increased more than 3-fold to almost 11 hours (74). Post-

transcriptional regulation of the renin gene was not well studied until the renin response

to forskolin in the lung carcinoma cell line Calu-6 was tested. Lang et al. observed that

12

transfected human renin promoter transcription reporter constructs were not significantly

induced upon forskolin treatment despite a robust response of the steady state mRNA

(75). Similar experiments from Florence Pinet’s lab largely supported this result (76).

Nuclear run-on analysis from Lang et al. indicated only a small increase in transcription

al activity after 1 hour of forskolin treatment, but none after 24 hours. They concluded

that stimulation of the cAMP pathway in these cells led primarily to increases in mRNA

stability rather than transcriptional activity. Sinn et al. went on to show that the increase

in half-life was so great that there was no decay in renin mRNA 60 hours following

pretreatment with forskolin (77). The difference in stability was not due to a different 5’

or 3’ UTR found in Calu-6 cells than that in the kidney mRNA. These results indicate

that the regulation of renin expression in these cells is almost entirely post-

transcriptional. Although not an optimal cell line for investigating renin transcription,

others have used it to identify RNA-binding proteins that regulate mRNA stability.

Using the Calu-6 cell line as a source of cytoplasmic protein, two groups have

attempted to determine what proteins might regulate cAMP-induced renin mRNA

stability (78, 79). As an initial screen, both considered the conservation of both the 5’

and 3’ UTRs of human renin mRNA. Because the 5’ UTR is poorly conserved and short

(32-44 nucleotides) it probably does not play a large role in mRNA stability. The 3’

UTR appears to be well conserved despite a lack of traditional AU-rich destabilizing

elements. However, there are some regions that resemble known RNA-binding protein

motifs from other genes. An atypical AU-rich element identified by Adams et al. appears

to potentially function as a destabilizing element and does not interfere with translation

(78). They go on to identify HuR, HADHB (hydroxyacyl-CoA dehydrogenase/3-

ketoacyl-CoA thiolase/enoyl-CoA hydratase trifunctional protein β-subunit), and CP1

(poly(C) binding protein-1) as 3’UTR binding proteins; and all three showed induction by

forskolin treatment of Calu-6 cells. One of these proteins, HuR, acts as a stabilizing

protein, whereas HADHB destabilizes renin mRNA. The HADHB protein is interesting

13

because of its specific localization in the JG apparatus. However, the role of HADHB in

JG cells has yet to be directly tested.

Skalweit et al. were able to identify six proteins that can bind to the renin mRNA

3’ UTR. These include: heterogeneous nuclear ribonucleoproteins hnRNP E1 and

hnRNP K (poly(C) binding proteins), dynamin, nucleolin, YB-1, and MINT-homologous

protein. Upon treatment of Calu-6 cells with forskolin, hnRNP E1, dynamin, nucleolin,

and YB-1 protein expression were all upregulated. Because all of the proteins identified

by these groups are induced by the cAMP pathway, it is difficult to know which of these,

if any, is participating in the stabilization of renin mRNA following activation. There is

no compelling evidence for increased 3’UTR binding after forskolin treatment in Calu-6

cells from either group. However, dynamin has been found to be downregulated by

increases in intracellular Ca2+ in As4.1 cells (64). Knockdown of dynamin in As4.1 cells

leads to a large decrease in steady state renin mRNA. Given the link between Ca2+ and

cAMP in renin expressing cells, it is reasonable to speculate that dynamin serves as the

link between the cAMP pathway and mRNA stability. However, it seems counter-

productive to control post-transcriptional regulation of one gene through transcriptional

regulation of another. The only related proteins identified by both groups are the

poly(C)-binding proteins hnRNP E1, hnRNP K, and CP-1. These proteins function in

mRNA stabilization, splicing, and translational regulation (80). There is therefore

potential for regulation by these proteins at other levels of post-transcriptional processing

such as splicing and translation.

Regulation of Renin Secretion

The same signals that stimulate or inhibit renin transcription also affect renin

secretion. Renin is first translated into preprorenin, containing a signal peptide which

directs the protein into the secretory apparatus. In the endoplasmic reticulum, the signal

sequence is removed. Prorenin then moves to the Golgi apparatus where it can be

14

glycosylated and sorted into the regulated pathway. The prosegment is then cleaved

inside the granule to form active renin. Constitutive and regulated pathways contribute to

the secretion of renin. The regulated pathway is controlled by a protein kinase A

dependent mechanism mediated by increases in cAMP (81). Those increases in cAMP

can be controlled by adenylate cyclase activation or phosphodiesterase inhibition (82).

As mentioned previously, increases in intracellular Ca2+ paradoxically inhibit renin

secretion. Renin and parathyroid hormone are the only two secreted proteins known to

be inhibited by increases in Ca2+. There is now evidence to suggest that Ca2+ inhibition

of renin secretion can be controlled by the calmodulin-calcineurin pathway which is

independent of PKA (83).

Sorting into the regulated pathway is dependent upon the presence of the

prosegment and a dibasic pair of amino acids at position 42-43 (84). This dibasic amino

acid pair seems to be the target of a processing enzyme important for renin’s sorting in

the secretory pathway. The prosegment does, however, contribute to the processing

controlled by the amino acid pair (85). It is the absence of the signal peptide and one-

third of the prosegment in the exon-1b and exon-1c renin isoforms that have led to the

hypothesis that these represent intracellular forms. Transient transfection of the exon-1b

cDNA into cells led to intracellular retention of active renin protein (86). The

glycosylation of renin is important for sorting as well. We know this because the Ren-2

protein lacks three asparagine residues that serve as glycosylation sites in Ren-1. In mice

with Ren-1 knocked out but Ren-2 preserved, JG cells completely lack secretory granules

(87). The phenotype was rescued only when Ren-1 knock out mice were backcrossed to

a BAC transgenic with Ren-1 and Ren-2, but not Ren-2 alone (88). For some reason, the

sorting of renin into the regulated pathway is rather inefficient because only ~25% of the

prorenin is directed into that pathway, while 75% is secreted without further processing

by the constitutive pathway (89).

15

Although understanding the renin secretory pathway is important for determining

how its release is controlled, it probably does not play a major role in how much is

released. The number of granules per JG cell does not vary under physiological

conditions that stimulate renin secretion (90). Long term regulation seems to favor

recruitment of more renin expressing cells rather than modification of granule

morphology or amount of release per cell (91-94). Although renin content per cell is not

altered, the level of renin mRNA per cell, the number of JG apparatuses, and the number

of cells expressing renin is increased (95). This recruitment of renin expressing cells is

an interesting phenomenon. As discussed in the next section, cells which express renin

during development apparently retain their ability to express renin and are recruited do so

under conditions when homeostasis is threatened. The constitutive pathway responsible

for prorenin release is tied to the transcriptional activity and post-transcriptional

regulation of the gene. During an acute stimulus the levels of active renin increase

whereas there is no change in prorenin (96). Thus, regulation of renin secretion is

controlled primarily by the regulated pathway. A long term stimulation results in an

increase in both prorenin and active renin indicating that the synthesis of renin (i.e. renin

expression) plays a more important role when renin secretion is chronically stimulated.

Development of the Renin Expressing Cell

Renin is absolutely essential for the normal development of the kidney. Renin

knockout mice die between birth and weaning due to severe renal defects and impaired

electrolyte retention. The survival of knockout mice must be rescued by saline injection

of neonates (37). These mice have no detectable angiotensin I or II, kidney fibrosis,

hydronephrosis, shrinkage of the tubules, and hypertrophy of the renal arterials. As

expected, they are hypotensive and unable to concentrate their urine. The phenotype of

these mice is similar to that of Potter’s syndrome (tubular dysgenesis) patients, a subset

of which have mutations in the renin gene that result in a lack of expression or inactive

16

protein (35). Mutations in other components of the RAS, including ACE,

angiotensinogen (AGT), and Ang II receptor (ATR) were also discovered. Patients with

mutations in these genes had renin expression in the mesangial cells and arteriolar

smooth muscle cells whereas it is normally restricted to the JG cells. This recapitulates a

phenomenon observed in animals treated with ACE inhibitors or ATR antagonists as well

as fetuses exposed to them (95, 97, 98). Indeed, one of the problems with pre-eclampsia,

a syndrome of hypertension during pregnancy, is that the patients cannot be treated with

ACE inhibitors or ARB, as these will affect renal development in the fetus. This has

been described as a recruitment or de-differentiation of cells that do not normally express

renin but retain the capacity to do so. Recruitment occurs due to a lack of negative

feedback, such as Ang II action, or presence of a positive stimulus like decreased

perfusion pressure in an attempt to maintain homeostasis. The pattern of expression

observed is akin to that seen during kidney development where renin is expressed

throughout the arterial tree of the kidney (99). As kidney development progresses, renin

expression becomes gradually restricted to the smaller vessels until it is localized to the

JG cells in the adult kidney. Recruitment of renin expressing cells under threatened

homeostasis reverses this pattern. What determines renin cell fate and what signals

control the recruitment of non-renin expressing cells has become an area of focus to

better understand the control of renin expression.

Renin expression can first be observed in the metanephric mesenchyme of the

embryonic kidney. Importantly, this is before kidney vascularization has occurred and

the renin cells do not yet express endothelial or smooth muscle cell (SMC) markers

(100). At embryonic day 16 (E16) renin cells acquire SMC, but not endothelial markers.

This indicates that renin cells are a precursor for SMCs in the kidney. However, they are

not the only source as there are SMCs in the metanephric kidney that do not express

renin. Lineage tracing reveals that renin expressing cells can give rise to smooth muscle

cells, extraglomerular and intraglomerular mesangial cells, cells of Bowman’s capsule,

17

and proximal tubule cells (101). The same cells labeled by lineage tracing but no longer

expressing renin stained positive for renin when mice were subjected to low sodium diet

plus ACE inhibition. Therefore, all cells that once expressed renin were capable of being

recruited under conditions of threatened homeostasis.

What molecular signals are responsible for the development of the renin

expressing cell and its restriction to the JG apparatus under normal conditions?

Intercellular communication mediated by gap junctions is playing a critical role.

Connexins 40, 43, and 37 are all expressed in JG cells (102). Another, connexin 45, is

restricted to the vascular smooth muscle cells in the adult kidney, but is co-expressed

with renin in cells during development. It has also been detected in the glomerulus and

mesangium (103). Connexin 40 expression is found in endothelial cells of the kidney

vasculature along with the intraglomerular and extraglomerular mesangium (102, 104).

Their localization may provide important information regarding their role in controlling

renin expression and localization.

Connexin 40 is necessary for the proper localization of renin expression in the

adult kidney. In connexin 40 global knockout mice (Cx40-KO) renin is found aberrantly

expressed along the afferent arteriole and in the extraglomerular mesangium (105).

Global knockout of connexin 40 results in hypertension and increased PRA as a result a

lack of negative regulation by increased perfusion pressure and AngII (106, 107). In fact,

isolated perfused kidneys from connexin 40 knockout mice show a paradoxical induction

in renin secretion rate when perfused with AngII. The lack of negative regulation

appears to be due to the absence of a response to extracellular calcium. This suggests

that the signal initiated by a reduction in extracellular calcium is not being transmitted to

the JG cells. The lack of a baroreceptor response is dependent on the presence of

connexin 40 in the JG cells rather than the endothelial cells (104). There is also a defect

in TGF inhibition of renal blood flow in Cx40-KO mice that is probably a reflection of its

loss in the extraglomerlular mesangium (108). Connexin 45 also appears to transmit a

18

negative regulatory signal as well. Knockout of connexin 45 in the vasculature of the

kidney results in an increase in renin and blood pressure (103). Vascular smooth muscle

cells from those knockout kidneys were found to transmit Ca2+ waves at a slower rate.

The role of connexin 43 is less well characterized, but it may propagate a positive

stimulus for renin secretion and expression in response to a decrease in perfusion

pressure (109). Lack of this connexin abrogated the increase in renin expression and

secretion in response to decreases in perfusion pressure. Again, Ca2+ appears to play an

integral role in renin expression and the recruitment of renin expressing cells. Connexins

may play an important role in transmission of calcium fluxes from cell to cell in response

to physiological cues.

Given the integral role of cAMP in the control of renin expression and secretion it

should come as no surprise that it is also involved in renin cell development and

recruitment. Using a dual fluorescent reporter mouse, Pentz et al. were able to isolate

cells of the renin cell lineage from renal arterial smooth muscle (110). Forskolin

induction of the cAMP pathway was capable of inducing renin expression and increased

histone acetylation around the promoter. Therefore, non-renin expressing cells of the

renin cell lineage maintain the ability to re-express renin upon stimulation of the cAMP

pathway. The histone acetylation is most likely mediated by CBP/p300 recruitment by

cAMP stimulated PKA phosphorylation of CREB and its subsequent binding to CREs in

the promoter and/or enhancer. In fact, CBP/p300 are absolutely essential for the

development of the renin expressing cell (111). Given the importance of Ca2+ in

controlling the production of cAMP in renin expressing cells it is not a stretch to assume

that the Ca2+ signals likely to be transmitted by connexins is modulating the recruitment

of non-renin expressing cells during threatened homeostasis.

19

Figure 1-1. Alignment of mouse and human enhancer.

Numbers indicate the position relative to the transcription start site. Mouse enhancer binding elements are labeled with their corresponding transcription factor indicated to the right. The HRE is boxed.

20

CHAPTER 2

CONTROL OF RENIN EXPRESSION BY NUCLEAR RECEPTORS

Introduction

Nuclear receptors are a specific class of the transcription factor family that share

a number of structural features. These features include (from N-terminus to C- terminus)

the transactivation domain (AF-1), a DNA binding domain (DBD), and a C-terminal

ligand binding domain (AF2). The AF-1 domain sequence varies the most between

nuclear receptors and sometimes serves as a recognition signal for co-activators and other

transcription factors. This domain can also serve as a site for post-translational

modifications that control nuclear receptor activity. The DBD contains two zinc finger

motifs that are involved in DNA binding but can also serve a role in dimerization with

other nuclear receptors and post-translational modifications. Nuclear localization signals

(NLS) can also be contained within the DBD. The AF2 domain serves as the ligand

binding site, binds co-regulators, serves as a site for dimerization with other nuclear

receptors, and contains NLSs. There are no known ligands of any kind for 20 of the 48

known human nuclear receptors which are therefore called “orphan” receptors. The other

nuclear receptors have either an endogenous or synthetic ligand that activates their

activity.

Nuclear receptors control cell responses by a variety of methods. Nuclear

receptors bind to DNA via variations of the RGGTCA sequence, also called the hormone

response elements (HREs). These response elements can form direct repeats, inverted

repeats, or everted repeats separated by different lengths of spacer. The nuclear receptors

can bind these elements as monomers, homodimers, or heterodimers. When bound to

DNA they can recruit co-regulators that can modify chromatin or other transcription

factors and interact with the general transcription machinery. However, their function

can be exerted through other mechanisms. Some exert their function independent of

21

direct DNA binding through transrepression. Others have been shown to have non-

genomic effects by interacting with signal transduction pathways (112-114).

The regulation of nuclear receptor function is not just controlled by the presence

or absence of ligand. Control of nuclear receptors themselves can be through the nature

of the element to which they bind or post-translational modifications like

phosphorylation, sumoylation, and ubiquitination. These then affect their interactions

with co-regulators or other transcription factors and/or DNA binding.

Because of their diverse control and function, nuclear receptors have been the

target of much research in transcriptional regulation and how those signals control

physiology. They also are attractive for drug development because of their ability to be

controlled by ligands. Orphan nuclear receptors are particularly interesting because they

control a variety of physiological functions yet lack any known ligand. Some may lack a

ligand altogether, but others may provide a novel link between physiological cues and

responses involved in disease.

Because of the presence of an HRE within the renin enhancer, our lab has been

interested in determining what nuclear receptors might regulate renin expression. We

have hypothesized that one of the nuclear receptors that bind to the HRE is responsible

for the robust induction of the promoter. Despite the identification of at least four nuclear

receptors (RAR, RXR, EAR2, and PPARγ) that can bind to the HRE and regulate renin

expression, none of them have the large impact on renin expression that would be

expected based on promoter activity assays. We have previously identified and

characterized Nr2f6 as a negative regulator of the renin promoter. Additionally, As4.1

cell nuclear extracts result in four HRE shift complexes in electrophoretic mobility shift

assays that cannot be completely accounted for by RAR, RXR, and Nr2f6 (115, 116).

Using the same yeast one-hybrid screen that identified Nr2f6, we identified another

orphan nuclear receptor, Nr2f2 (Arp-1, Coup-TFII). This receptor is a member of the

same group of nuclear receptors (subfamily 2) and is thus closely related to Nr2f6.

22

Because Nr2f6 (EAR-2) is a negative regulator of the renin promoter, we hypothesized

that Nr2f2 (Coup-TFII) would be as well. We also wanted to determine the role of Nr4a1

(Nur77) and Nr4a2 (Nurr1), which were found to be expressed in As4.1 cells and

identified in a BLAST search of orphan nuclear receptors to have homology with Nr2f6.

Methods

RT-PCR

As4.1 total RNA was isolated using the PurelinkTM RNA Mini Kit. After RNA

isolation, 1 µg was reverse transcribed using 200 units of Superscript III in a 20 µl

reaction (Invitrogen). Reactions were incubated at 50°C for 30 minutes, 55°C for 15

minutes, 60°C for 15 minutes, and 70°C for 15 minutes to inactivate the reaction.

Orphan receptor cDNAs were amplified using AmpliTaq Gold (Applied Biosytems) with

1X PCR buffer containing 1.5 mM MgCl2. See Table 3-1 for primer sequences.

cDNA Expression Plasmids

Expression plasmids were constructed using cDNAs for Nr2f2, Nr4a1, and Nr4a2.

The cDNAs were obtained by reverse transcription of As4.1 totRNA isolated using the

PurelinkTM RNA Mini Kit. They were PCR amplified using HiFi Platinum Taq

Polymerase (Invitrogen) using primers designed at the translation start and termination

codons. Amplified cDNAs were TOPO cloned into pCR2.1 (Invitrogen) and

subsequently clone into pcDNA3.1(+) (Invitrogen).

Luciferase Assay

As4.1 cells (ATCC CRL2193) were split and transfected the following day using

a 3:1 ratio of Fugene 6 (Roche) to plasmid DNA. One microgram of m4.1kP-luc (117)

and 1 ng phRL-TK (Promega) were included in all transfections. Each well was

transfected with 1 µg cDNA or shRNA expression plasmid corresponding to the specific

23

nuclear receptor. Forty-eight hours post-transfection cells were lysed and extracts

analyzed using Promega’s Dual-Luciferase Assay System.

Orphan Nuclear Receptor Knockdown

Adenoviruses expressing shRNA to GFP, Nr2f2, and Nr2f6 were constructed and

tested as previously described (118, 119). As4.1 cells were infected at 60% confluency

using an MOI of 100, 24 hours after being split into 6-well dishes. Adenovirus and

polybrene (5µg/ml; Millipore) were mixed in serum free DMEM and added to duplicate

wells for each shRNA. After a six hour incubation, cells were washed and complete

DMEM (10% FBS, 100 U/ml penicillin, 100 mg/ml streptomycin) was added. Forty-

eight hours later, total RNA was isolated using the PurelinkTM RNA Mini Kit and protein

extracted using RIPA buffer (50 mM Tris-Cl pH 7.4, 150 mM NaCl, 1% NP-40, 0.5%

Sodium deoxycholate, 0.1% SDS).

The cDNA was obtained as indicate above, diluted 1:20, and gene expression was

measured using Taqman® Gene Expression Master Mix (Applied Biosystems) and

Taqman® probes or iQ SYBR Green Supermix (Bio-Rad) and primers. Nr2f2, Nr2f6, and

Nr4a1 probes were from Applied Biosystems as listed on the Nuclear Receptor Signaling

Atlas website (www.nursa.org, Table 3-1). The renin (Mm02342888_gH) and β-actin

probes (4352933E) were from Applied Biosystems. Data was analyzed using the 2-∆∆Ct

method to calculate fold changes relative to shGFP samples. Assay PCR efficiency was

determined to be 90-105% using a 7-log serial dilution series of the cDNA samples.

Protein extracts were quantified and 10 µg of each were mixed with SDS sample

buffer. Samples were loaded and run on 10% SDS-PAGE gels, transferred to PVDF

(Millipore), and probed for renin (Santa Cruz; sc-22671), Nr2f2 (Perseus Proteomics; PP-

H7147-00), and actin (Abcam; ab1801). Horseradish peroxidase conjugated secondary

antibodies included the Goat TrueBlot® (eBioscience; 18-8814) for renin blots, the ECL

http://www.nursa.org/

24

anti-Mouse IgG (GE Healthcare; NA931) for Nr2f2 blots, and the ECL anti-Rabbit IgG

(GE Healthcare; NA-934) for actin blots.

Electrophoretic Mobility Shift Assay and Supershift Assay

Electrophoretic mobility shift assays (EMSA) were carried out using double

stranded DNA probes corresponding to the HRE were designed with 5’-GATC overhangs

and labeled using [α-32P]dATP. The wild-type HRE sequence is 5’-

GATCTGGTGACCTGGCTGTACTCTGACCTCTCAGAT-3’ and mutant probes µb:

5’-GATCTGGTGACCTGGCTGTACTCTTTCCTCTCAGAT-3’, µc: 5’-GATCTGGTT

TCCTGGCTGTACTCTGACCTCTCAGAT-3’, µbc: 5’-GATCTGGTTTCCTGGCTGT

ACTCTTTCCTCTCAGAT-3’. In vitro translated proteins were generated using the

TNT® Quick Coupled Transcription/Translation System (Promega). Parallel reactions to

assess protein production were run in which proteins were labelled using 35S-Methionine.

Probes were incubated at room temperature for 30 minutes with 1 µl of unlabeled

in vitro translated protein or 6 µg of As4.1 nuclear extract in Tris binding buffer (10 mM

Tris-Cl pH 7.4, 1 mM EDTA pH 8.0, 60 mM KCl, 10 mM DTT, 0.1% Triton X-100, 4%

Glycerol) with 1 µg poly[d(I-C)]. Binding reactions were loaded onto 5% native

polyacrylamide gels and run for 2 hours. Gels were dried, exposed to phospho-screens

overnight, and scanned using a Molecular Dynamics Storm 840. Supershift analysis was

performed by adding 1 µg of the appropriate antibody after the initial incubation period

for 15 minutes on ice prior to electrophoresis.

DNA Affinity Purification Assay

DNA affinity purification assays were carried out with slight modifications as

described by Mittler et al. using Biotin-TEG 5’-labeled double-stranded DNA probes

(120, 121). The sequence for the WT probe was 5’-CAAAACTGCAGGATGGTGACC

TGGCTGTACTCTGACCTCTCAGAT-3’ and for the mutant probe was 5’- CAAAACT

GCAGGATGGTTTCCTGGCTGTACTCTTTCCTCTCAGAT-3’. Nuclear extracts from

25

As4.1 cells (40 µg) were mixed with 80 pmol of double stranded probe in the same

binding buffer as that used in EMSAs with protease and phosphatase inhibitors (Roche),

plus 4 µg poly[d(I-C)] (Roche) for a total binding reaction of 40 µl. Nuclear extract and

probe were incubated on ice for 30 minutes followed by addition of 50 µl of streptavidin

conjugated Dynabeads® MyOne™ C1 (Invitrogen). Following a 90 minute incubation at

4°C while rotating, beads were collected using the DynaMag™-2 magnet (Invitrogen)

and washed three times with binding buffer. Beads were subsequently boiled, separated,

and extracts loaded onto a 10% SDS-PAGE gel. Western blots were probed for Nr2f2

and Nr2f6 (Abcam, ab65012).

Chromatin Immunoprecipitation

Chromatin immunoprecipitation (ChIP) was performed as described in the EZ-

ChIP (Millipore) kit manual. Briefly, chromatin extracts from As4.1 cells were subjected

to sonication using a Fisher Scientific Sonic Dismembrator Model 500 at an amplitude of

40% for 10-15 cycles of 10 seconds on and 10 seconds between each pulse. Sheared

chromatin was subjected to immunoprecipitation using 5 µg of the antibodies indicated

previously along with 1 µg of IgG provided with the kit. Immunoprecipitated chromatin

was subjected to PCR for 30 cycles using primers flanking the HRE (5’- TTGGACCCTC

TCCATTCCTTCACG-3’, 5’- ATGCGCTATCACAACCAGCCACTC-3’) and a region

in intron 5 (5’- ATTTGAGGGTGGGAAGGAAGG-3’, 5’- ATGAACTGGAAGAGGAC

CGAG-3’) of the renin gene.

Immunofluorescence

As4.1 cells were grown to 80% confluency in 24-well dishes with 12 mm #1

coverslips. Cells were washed with PBS and fixed for five minutes in PBS with 1%

Triton X-100 and 3.7% formaldehyde. They were subsequently washed three times in

PBS with 0.1% BSA. Cells were permeabilized and blocked for 1 hour at room

temperature in blocking buffer (1X PBS, 1% BSA, 5% goat serum, 0.2% Tween 20, 0.2%

26

NP-40) with slight agitation. Coverslips were washed and incubated with primary

antibody (1:100 dilution α-Nr2f2 and α-Nr2f6 and 1:50 dilution Nr4a1 (Santa Cruz; sc-

7014)) in blocking buffer overnight at 4°C. After washes, cells were incubated for 1 hour

with secondary antibody (goat anti-rabbit Alexa Fluor 586 or anti-mouse Alexa Fluor

488; 1:200 dilution) in blocking buffer. Cells were washed and mounted with

Vectashield with DAPI or incubated with 10 µM TOPRO3 for 15 minutes. Coverslips

incubated with TOPRO3 were washed and mounted with Vectashield for fluorescence.

Cells were imaged and observed using a Nikon Eclipse E600 fluorescent scope equipped

with a SPOT advanced digital camera (Diagnostic Instruments, Inc.)

Results

Initially, the HRE was shown to be a response element that RAR/RXR could bind

to and mediate the induction of renin expression by retinoic acid. We now recognize it as

an element that can bind several other nuclear receptors that regulate the renin promoter.

Previously, we identified Nr2f2 and Nr2f6 as HRE binding proteins using the HRE as

bait in a yeast one-hybrid screen. Nr2f6 was further characterized to act as a negative

regulator of the renin promoter via binding to the HRE in the enhancer. However, Nr2f6

only accounted for one of at least four proteins that could complex with the HRE and the

role of Nr2f2 was not determined. Here, in order to establish the role of Nr2f2 in

regulating renin expression, we are using Nr2f6 as a positive control.

By searching As4.1 microarray data obtained previously in our lab by Hana Itani

and performing RT-PCR we have identified several nuclear receptors that are expressed.

All of the nuclear receptors tested in my studies were detected via microarray and RT-

PCR (Table 2-2 and Figure 2-1A). Immunofluorescence confirmed the expression of all

three nuclear receptors. Localization of Nr2f2 and Nr4a1 were almost completely

nuclear, while Nr2f6 showed cytoplasmic and nuclear localization (Figure 2-2). The dual

localization of Nr2f6 suggests that it may shuttle between the cytoplasm and nucleus.

27

Quantitative PCR shows that Nr2f2 had the lowest expression level of the three nuclear

receptors tested. The expression level of Nr2f6 was two-fold greater than Nr2f2 and

Nr4a1 was five-fold greater (Figure 2-1B). Thus, several nuclear receptors are found in

As4.1 cells and of the three tested in this study, Nr4a1 has the highest expression level.

In order to test whether Nr2f2 can regulate the mouse renin promoter we

cotransfected cDNA expression vectors for Nr2f2 and Nr2f6 with m4.1kP-luc reporter

vector into As4.1 cells. The m4.1kP-luc vector contains 4.1 kb of the 5’ upstream

sequence of the mouse Ren1 gene driving expression of firefly luciferase.

Overexpression of Nr2f2 led to a 70% reduction in promoter activity compared to the

empty vector control (Figure 2-3A). The overexpression of Nr2f6 and Nr4a1 resulted in

an approximately 44% and 60% reduction, respectively. Another orphan nuclear

receptor, Nr4a2, had no effect on promoter activity. Because overexpression of these

nuclear receptors could force interactions with other proteins or cis-acting elements that

do not normally occur, we performed the reverse experiment where we co-transfected

shRNAs targeting Nr2f2 and Nr2f6 with m4.1kP-luc. Knockdown of Nr2f2 and Nr2f6

produced a greater than 2-fold increase in promoter activity (Figure 2-3B). When Nr4a1

was knocked down it resulted in a surprising decrease in promoter activity. However,

this did not reach statistical significance. The data here suggest that Nr2f2 has the

potential to negatively regulate the mouse renin promoter and supports our previous data

for Nr2f6. In contrast, Nr4a1 exerts opposite effects when overexpressed or knocked

down.

We next used EMSAs to determine if Nr2f2 could directly bind to the HRE.

When in vitro translated proteins were combined with 32P-labeled HRE probe and

DR1(G) or NBRE control probe, Nr2f2 and Nr2f6 show clear binding to the HRE that is

effectively competed away by excess WT cold probe (Figure 2-4A and 2-4B). In

contrast, Nr4a1 shows no binding to the HRE but does bind the control probe (Figure 2-

4C). We combined equivalent volumes of in vitro translated Nr2f2 and Nr2f6 extracts

28

with the HRE probe to determine if they could heterodimerize to form different shift

complexes. Two specific shift complexes are formed that correspond to the individual

complexes formed when Nr2f2 and Nr2f6 are incubated with the HRE probe alone

(Figure 2-5A). Each complex is effectively competed away by 100 fold excess wild-type

(WT) cold probe. That competition is lost when either half-site (µb or µc) or both (µbc)

are mutated. This suggests that both half-sites are necessary for binding to the HRE and

therefore Nr2f2 and Nr2f6 bind as dimers. This is consistent with the proposed

functional DNA binding form of Nr2f2 (122, 123). However, this does not eliminate the

possibility that Nr2f2 and Nr2f6 cannot bind as monomers, because some competition is

maintained with an intact b site and mutated c site. Therefore, binding to the HRE may

be primarily dependent on a single intact b site suggesting that Nr2f2 and Nr2f6 are

binding as monomers. It has been shown previously that Nr2f2 and Nr2f6 can form DNA

binding heterodimers (124). The top complex was entirely supershifted by the addition

of Nr2f2 antibody, whereas the bottom complex was shifted by Nr2f6 antibody. Neither

antibody supershifted both bands, indicating that Nr2f2 and Nr2f6 do not form

heterodimers when binding to the HRE. However, this does not eliminate the possibility

that they can heterodimerize with other nuclear receptors like RXR. Altogether, these

data indicate that the negative regulation on the renin promoter can be mediated by direct

binding of Nr2f2 and Nr2f6 to the HRE. In contrast, Nr4a1 does not appear to mediate

its effects though direct binding to the HRE. If Nr4a1 can bind to the HRE, it is through

binding to another transcription factor.

To verify that Nr2f2 is one of the proteins from As4.1 cells that forms a complex

with the HRE, we performed the same EMSA analysis with nuclear extracts. As

observed previously, four shift complexes (a, b, c, d) were formed that are effectively

competed away by excess WT probe (Figure 2-5B) (116). Complexes b and c are still

competed away when only one of the half-sites is mutated (µb or µc), but is lost with

mutation of both half-sites (µbc). In contrast, mutant half-site probes are less effective

29

competitors for unidentified complexes a and d. We hypothesized that complexes b and c

correspond to Nr2f2 and Nr2f6 based on their similarity to the shift complexes formed by

in vitro translated Nr2f2 and Nr2f6. In support of complex c being Nr2f6, the addition of

Nr2f6 antibody resulted in the supershift of complex c and replicates our previous results

(116). However, none of the complexes was supershifted or reduced by addition of

Nr2f2 antibody. Therefore, we performed DNA affinity puridication assays (DAPA) as

an alternative for identifying Nr2f2. Both Nr2f2 and Nr2f6 show clear enrichment for

binding to the WT DAPA probe versus a mutant probe with both half-sites mutated

(Figure 2-5C). Thus, Nr2f2 and Nr2f6 can directly bind to the HRE and may form two of

the four complexes from EMSA.

Whether or not Nr2f2 and Nr2f6 can bind to the enhancer in its native chromatin

context was addressed using ChIP. Both nuclear receptors showed clear enrichment over

IgG immunoprecipitated chromatin. As a positive control, Creb1 antibody was used to

precipitate chromatin and showed enrichment as well. No signal was detected when a

region approximately 10.5 kb downstream of the enhancer in intron 6 was probed,

indicating that the signal was specific for chromatin around the HRE. This indicates that

both Nr2f2 and Nr2f6 can bind to chromatin around the HRE to control renin expression.

Luciferase assays, EMSA, and DAPA all support a mechanism by which Nr2f2

negatively regulates renin expression through its binding to the HRE. We aimed to

substantiate that role further by determining the effect of Nr2f2 knockdown on the

endogenous renin gene in its native genomic and chromatin context. Using the same

shRNAs as used in luciferase assays, we specifically knocked down Nr2f2, Nr2f6, and

Nr4a1 in As4.1 cells and measured renin mRNA levels using qPCR (Figure 2-6A).

Knockdown of Nr2f2 had no effect on baseline renin expression despite a robust

reduction of Nr2f2 mRNA. In contrast, Nr2f6 knockdown resulted in an approximately

1.8-fold increase in renin. The lack of effect on renin expression was not due to the

absence of Nr2f2 protein knockdown, because that mirrored the reduction of mRNA

30

(Figure 2-6B). This would indicate that although Nr2f2 can negatively modulate renin

promoter activity it is not able to function in that manner when the renin gene is in its

native chromatin context. Surprisingly, renin expression decreased ~2-fold when Nr4a1

was knocked down (Figure 2-6A). That would support the result seen in luciferase

experiments when Nr4a1 was knocked down, suggesting that it is a positive regulator of

renin expression.

In an attempt to further validate nuclear receptor knockdown and the specificity of

its effect on renin expression, we analyzed the expression levels of other potential target

genes. These potential targets were identified from a genome-wide scan of Nr2f1 binding

sites done previously (125). Nr2f1 is the other member of nuclear receptor subfamily 2

group F and shares a high degree of homology with Nr2f2. That analysis formed a

consensus Nr2f1 binding motif based on every binding site sequence published at that

time. The consensus sequence that was used for scanning the genome was very similar to

the TGACCT repeat motif of the enhancer. Genes that were determined to be

differentially expressed and have a ChIP validated binding site for Nr2f1 were used. Of

the genes tested using qPCR, superoxided dismutase (Sod1) was upregulated to a similar

level of that seen with renin when Nr2f6 was knocked down (Figure 2-7). Unexpectedly,

cellular retinoic acid binding protein 1 (Crabp1) was downregulated with knock down of

Nr4a1. This corroborates further that knock down of these nuclear receptors is sufficient

to change gene expression. However, the effect is not specific to the renin gene. Both

genes are involved in pathways that regulate renin. Retinoic acid stimulates renin

expression and hydrogen peroxide that can be produced by Sod1 is a negative regulator.

This suggests that renin expression may be controlled by Nr2f6 and Nr4a1 through direct

and indirect mechanisms. Unfortunately, we have not yet identified a gene whose

expression changes following knockdown of Nr2f2.

31

Discussion

Nuclear receptors are a diverse family of transcription factors that are regulators

of many physiological processes. They have been found to regulate development,

metabolism, vascular function, circadian rhythm, and reproduction. Many are activated

by ligands that allow them to respond to physiological changes in the body and modify

transcriptional programs. The nuclear receptor superfamily is an attractive

pharmacological target because receptor-selective, cell-type selective, activity selective,

as well as partial, full, and inverse agonists have been developed (126). Some nuclear

receptors are identified as orphans because they lack a known endogenous ligand. These

have become increasingly interesting given the synthetic ligands that have been

discovered for orphan receptors such as Pparγ. It serves as a good example of activity

selective ligands for a nuclear receptor. In the case of Pparγ, the ligand MRL24 shows

moderate affects on Pparγ transcriptional activity, but strong inhibition of its

phosphorylation (127-129). These two pathways result in different gene expression

regulation (128, 129). Because of their pharmacological and therapeutic potential,

orphan nuclear receptors may provide novel targets for the treatment of hypertension.

We have previously identified the orphan nuclear receptor Nr2f6 (EAR2) as a

negative regulator of the renin gene. This led us to ask what other orphan receptors

might bind to the HRE of the renin enhancer and regulate promoter activity. Despite the

discovery of nuclear receptors that can regulate the renin promoter, none of them result in

the robust change that results when the HRE is mutated. Futhermore, we have yet to

identify all of the proteins in As4.1 cells that are able to bind to the HRE. Therefore, we

proceeded to characterize the role of Nr2f2 and Nr4a1 in regulating renin expression.

As a first line of study we utilized transient transfections of a vector containing

the mouse renin promoter driving expression of luciferase co-transfected with

overexpression or shRNA plasmids. Luciferase assays suggest that Nr2f2 and Nr2f6 act

as repressors of renin promoter activity. Nr4a1 showed conflicting results when

32

overexpressed or knocked down as both conditions showed a downregulation of promoter

activity. Nr2f2 and Nr2f6 can directly bind the HRE as indicated in EMSA and DAPA

experiments, but Nr4a1 does not. Luciferase and binding experiments taken together

suggest that the repressor activity of Nr2f2 and Nr2f6 are mediated through the HRE.

However, Nr2f2 and Nr2f6 appear to have divergent effects on baseline endogenous gene

expression. Nr2f2 knockdown did not affect renin expression, whereas Nr2f6

knockdown resulted in an increase. Interestingly, Nr4a1 knockdown resulted in a

decrease in renin expression indicating that it activates renin expression.

The main question that needs to be answered is why Nr2f2 had effects on

transfected promoters but not the endogenous. It is a member of the same subfamily as

Nr2f6 albeit more distantly than the other family member, Nr2f1 (Coup-TFI).

Compensation of Nr2f2 loss by Nr2f1 is unlikely because that would have been observed

in transfection experiments. It will be of interest to investigate the role of Nr2f1 in

regulation of the endogenous renin promoter to examine if the lack of regulation is

common to the two closer related family members. Nr2f6 was in vitro translated at a

much lower efficiency than Nr2f2 and thus smaller amounts were present in the binding

reactions in EMSAs. Despite being at a lower concentration, it showed similar binding

intensity to Nr2f2. The absence of Nr2f2 regulation might be related to the different

binding affinities for the HRE.

The argument could be made that Nr2f2 does not bind to the HRE in the native

chromatin context, but ChIP studies contradict that argument. There is a possibility that

binding detected in the ChIP experiments is at another site in the enhancer. The binding

of Nr2f2 to the early growth response 1 gene (Egr1) promoter is through indirect binding

to Sp1 (130). Therefore, binding to the enhancer might be through the Sp1 site upstream

of the HRE. If that is the case, loss of Sp1 or an additional factor may be required to

unmask a response at baseline. It is also possible that Nr2f2 does not modify baseline

activity but controls induction or repression by a signal that regulates renin. What that

33

signal might be will require further studies examining renin expression responses to

cAMP-PKA pathway stimulation or increases in intracellular Ca2+ with Nr2f2

knockdown. Examination of the crystal structure of Nr2f2 suggests that it is an auto-

repressed conformation but promotes transcription activity in multiple cell lines (122).

Furthermore, mutation of sites responsible for cofactor binding, dimerization, and ligand

binding reduce that transcription activity. These results suggest that there is a ligand for

Nr2f2 that can activate it. It is possible that Nr2f2 is regulated post-translationally, but to

date no modifications have been identified. Despite our finding that Nr2f2 does not

regulate baseline renin expression, it still deserves further attention. Interestingly, genetic

data in humans and Dahl salt-sensitive rats show an association of Nr2f2 with

hypertension (131-134).

My studies further supported the role of Nr2f6 as a negative regulator of renin

expression through binding to the HRE. I replicated both luciferase experiments and

showed that its knockdown resulted in an increase in renin expression. Although this

response is modest, inducing renin gene expression in As4.1 cells has been shown to be

difficult. Forskolin stimulation of renin expression in As4.1 cells requires

phosphodiesterase inhibition. Therefore, renin expression may be nearly maximally

stimulated in this cell line. Nr2f6 knockdown may result in a more robust increase when

the cAMP-PKA pathway is inhibited.

The mechanisms underlying Nr2f6 regulation and the genes which it regulates are

poorly understood. A recent study showed that Nr2f6 binding is inhibited when

phosphorylated by protein kinase C (PKC) (135). Although increases in calcium, which

can activate PKC, play an important role in negatively regulating renin expression, the

phosphorylation of Nr2f6 in As4.1 cells would prevent its binding and negative

regulation of the renin promoter. In that same study, in vitro kinase assays revealed a

phosphorylation of Nr2f6 by PKA. A mechanism by which PKA phosphorylation of

Nr2f6 leads to a decrease in its binding to the HRE and activation of renin expression is

34

an attractive one. In fact, previous studies in our lab show a robust inhibition of

endogenous renin expression when Nr2f6 is overexpressed (116). In that situation Nr2f6

levels may be high enough to provide an unphosphorylated pool capable of binding the

HRE and repressing the renin promoter.

The potential role of Nr4a1 as an activator of renin expression is of particular

interest. Although luciferase assays revealed divergent effects when Nr4a1 was

overexpressed or knocked down, it could be due to the nature of the experiments.

Overexpression might cause Nr4a1 to act as a sink by forming heterodimers with

transcriptional activators it does not normally bind with. In contrast, knock down

decreases its occupancy somewhere on the 5’ upstream sequence. The shRNA

experiments using luciferase assays support the notion that Nr4a1 is a positive regulator

of renin expression. Even though it does not directly bind to the HRE, it remains to be

determined whether it can bind to the enhancer indirectly. Control of renin expression by

Nr4a1 may still be mediated through the HRE by a transactivation pathway. Conversely,

it may bind to an unidentified motif elsewhere in the 5’ upstream sequence of the renin

gene.

Members of nuclear receptor subfamily 4 are unlikely to have endogenous ligands

due to their small binding pocket. However, regulation of their expression and

phosphorylation are important for their function. Their expression is induced by the

cAMP pathway that is so important in renin expression. Furthermore, Ken Gross and

Ariel Gomez have found Nr4a1 to be highly enriched in JG cells (personal

communication). However, my data from As4.1 cells is difficult to interpret given the

higher levels of cell death observed when Nr4a1 is knocked down. This may be due to

the nature of As4.1 cells. The Ren-2 promoter drives expression of T-antigen in these

cells which may be critical for their survival in culture. Thus, removal of an activator of

the renin promoter may result in a loss of T-antigen, loss of cell growth and ultimately

35

cell death. The effect of Nr4a1 does appear to be specific to the renin promoter since

levels of other genes tested in qPCR remain unchanged.

Further attention should be given to Nr4a1, but experiments should be conducted

in another cell line. The renin cell lineage cell line recently isolated by Ariel Gomez’s

lab may be a good model in which to conduct those experiments (129). These cells were

isolated by creating two separate reporter gene mice. One line had Cre recombinase

knocked into the endogenous Ren1d locus and a renin promoter transgene driving

expression of yellow fluorescent protein (YFP). This mouse was crossed with one that

had CFP knocked into the ROSA26 locus. Offspring positive for both transgenes would

express YFP in cells currently expressing renin and any cells that had expressed renin at

one time were marked with CFP. Therefore, cells of the renin lineage were marked with

CFP. The cells were then isolated by FACS sorting. These cells do not expresss renin at

baseline, but maintain their ability to do so upon cAMP induction. Therefore, they are

attractive for studying what signals are responsible for inducing renin expression.

Because Nr4a1 expression is induced by the cAMP pathway and it is enriched in JG cells,

it may participate in activating renin expression in non-renin expressing cells of the renin

lineage and maintenance of renin expressing cells.

In the process of validating knockdown of our nuclear receptors we identified two

other genes that were differentially regulated. By chance, these genes are involved in

pathways known to regulate renin expression. Hydrogen peroxide (H2O2) was found

previously in our lab to negatively regulate renin expression (136). Superoxide

dismutase 1, which was upregulated when Nr2f6 was knocked down, converts superoxide

radicals to oxygen and H2O2. The upregulation of Sod1 would presumably lead to

increases in H2O2 and a decrease in renin expression. This response is contradictory to

what might be expected since Nr2f6 led to an increase in renin expression. Of course, we

do not have any data suggesting that these cells actively generate superoxide, the

precursor to H2O2. In the case of Nr4a1 knockdown, Crabp1 was down regulated. It

36

binds to retinoids and decreases cell responses to them (137, 138). Again, this is

paradoxical since Nr4a1 and retinoic acid positively regulate renin expression. The gene

expression changes controlled in these cases might serve as negative feedback

mechanisms to control renin expression. Experiments specifically targeting Sod1 and

Crabp1 expression independent of Nr2f6 or Nr4a1 knockdown are needed to determine if

these changes do in fact modulate renin expression.

As a control in the ChIP studies, we employed an antibody to Essra. We did this

because of a recent report suggesting that Essra bound to the enhancer and regulated

renin expression. Although we confirmed the binding of Essra to the enhancer, we were

unable to replicate their effects on endogenous renin expression despite reasonable

knockdown of Essra.

My experiments have revealed that orphan receptors Nr2f2, Nr2f6, and Nr4a1 do

play a role in the control of renin expression. However, Nr2f2 and Nr2f6 are negative

regulators of the renin promoter, whereas Nr4a1 does not bind to the HRE. We initially

hypothesized that HRE binding proteins would be positive regulators given their potency

in activating promoter activity both in vitro and in vivo. Retinoic acid receptor exerts a

positive effect, but it is not very robust and may therefore play a minor role. There are

still two binding complexes from the EMSA analysis that remain unidentified. Whether

one of those complexes is RAR remains unexplored. If it is, that leaves at least one

complex to be identified. Alternatively, other transcription factors not detected by EMSA

could be binding to the HRE. These factors could have modest affinity for the HRE and

dissociate during electrophoresis. Furthermore, underrepresented factors may not be at

sufficient levels to compete with other factors for HRE binding despite the probe being

present in excess in the binding reaction. Future experiments will utilize our DAPA

protocol combined with mass spectrometry analysis. This will provide for a high

throughput unbiased approach for identifying HRE binding proteins from As4.1 cells.

This has been used previously to identify binding proteins for other motifs (120, 121).

37

Table 2-1. Primers and probes for gene expression analysis.

38

Figure 2-1. Expression of orphan nuclear receptors in As4.1 cells.

A) RT-PCR of As4.1 mRNA B) Quantitative PCR for the orphan nuclear receptors tested

39

Table 2-2. Summary of data from RT-PCR and microarray analysis of As4.1 mRNA

40

Figure 2-2. Immunofluorescence for Nr2f2, Nr2f6, and Nr4a1.

41

Figure 2-3. Luciferase activity of 4.1kP-luc when Nr2f2, Nr2f6, or Nr4a1 are overexpressed or knocked down.

A) Nr2f2 or Nr2f6 cDNA expressing plasmids co-transfected with 4.1 kP-luc. B) Plasmids expressing shRNAs to Nr2f2 or Nr2f6 co-transfected with 4.1kP-luc. Ratio of firefly luciferase (4.1kP-luc) to renilla luciferase (phRL-TK) plotted (N=5, * = p<0.05, One-way Repeated Measures ANOVA).

42

Figure 2-4. EMSA analysis of in vitro translated Nr2f2, Nr2f6, and Nr4a1.

Binding reactions were carried out with programmed extracts for empty vector (3.1), Nr2f2 (F2), Nr2f6, or Nr4a1. Different 32P-labeled probes for the hormone response element (HRE), control probes DR1(G) or NBRE, were incubated with lysates. Competition was performed by the addition of unlabeled (C) wild type (WT), mutated b site (µb), mutated c site (µc), or mutated b and c site (µbc) probes. Sequences of probes are displayed. Half-sties are bolded and mutated nucleotides are lower case.

43

Figure 2-5. Characterization of Nr2f2, Nr2f6, and As4.1 nuclear proteins to the HRE.

A) EMSA analysis using equivalent volumes of in vitro transcribed/translated Nr2f2 (F2)Nr2f6 (F6) or both (B) added to the same binding reaction with 32P-labeled d

, sDNA HRE

probe. Probes used for competition include the unmutated wild-type probe (WT) and probes with mutated half sites (µb or µc) or with both mutated (µbc). Supershifts were done by adding Creb1 (C1), Nr2f2 (F2), or Nr2f6 (F6) antibody to binding reactions. B,

a Biotin-TEG-labeled HRE probe and As4.1 cell nuclear extracts. Input= As4.1 nuclear extract, FT= flow through, WT= wild-type probe, M= mutant probe D) Chip assays probing the enhancer region or a region of intron 5. Precipitation was done using Nr2f2, Nr2f6, Creb1, or Esrra (Ea) antibody.

EMSA using As4.1 cell nuclear extracts. C) Western blots from DAPA assays using

44

Figure 2-6. Renin, Nr2f2, Nr2f6, and Nr4a1 expression following orphan receptor knock down in As4.1 cells.

) Renin, Nr2f2, Nr2f6, or Nr4a1 mRNA levels in As4.1 cells expressing shRNA to r2f2, Nr2f6, or Nr4a1 relative to mRNA in shGFP expressing cells. Expression levels

of mRNA was determined by RT-qPCR (N=6, *= p<0.05, One-way Repeated Measures ANOVA) B) Western blots for Nr2f2 using protein extracts from shGFP, shNr2f2, and shNr2f6 expressing As4.1 cells.

AN

45

Figure 2-7. Expression of other potential target genes following orphan receptor knock down.

Expression of Sod1, Crabp1, or Foxo3a in As4.1 cells following knockdown of Nr2f2, Nr2f6, or Nr4a1. Expression levels of mRNA was determined by RT-qPCR (N=6, *= p<0.05, One-way Repeated Measures ANOVA)

46

CHAPTER 3

CONTROL OF THE RENIN LOCUS BY CTCF

Introduction

The control of gene expression can be regulated through the interactions of

regulatory elements that reside large distances apart in the genome. There are

documented cases of interchromosomal contacts and intrachromosomal loops such as

those that occur during olfactory receptor and allele specific expression (139, 140). It is

therefore important to protect loci from the influences of other gene’s regulatory elements

to maintain their normal tissue and cell-specific gene expression. Insulators are

important regulatory elements that carry out this function. They are typically defined by

their ability to block communication between regulatory elements (enhancer blocking) or

block the spread of heterochromatin (barrier activity). A DNA binding protein found at

insulators, CCCTC-binding factor (CTCF), can mediate enhancer blocking activity

through a variety of mechanisms. The first correlation between CTCF and enhancer

blocking activity was shown for hypersensitive site 5 (HS5) of the β-globin locus.

Barrier activity however does not appear to require CTCF, thus it is primarily functional

in enhancer blocking insulators (141).

In our studies of the human renin kidney enhancer (KE) we have utilized large

genomic PAC transgenesis. These constructs are advantageous because they presumably

contain all of the necessary regulatory elements and provide protection against transgene

insertion site position effects of neighboring genes. This is critical given the difficulties

encountered in creating transgenic mice to tease out the regulatory elements important in

the human enhancer. Previous attempts have resulted in the ubiquitous expression or

variable regulation in response to physiological cues (142-146). This is probably due to

the influence of regulatory elements in genes around the site of insertion. In contrast,

larger constructs containing 25 kb, 35 kb, or 70 kb of 5’ upstream sequence show

47

appropriate cell- and tissue-specific expression and responses to all physiological input

(146, 147). Furthermore, the development of BAC recombineering has made these large

constructs amenable to the deletion and insertion of sequences within them. In fact, our

lab has used this method to delete both the KE and chorionic enhancer (CE) (146-149).

The PAC construct used in our lab contains 160 kb of human chromosome 1 with

the human renin gene (hREN), the upstream genes GOLT1A and KISS1, and the

downstream gene ETNK2. During the production and validation of PAC transgenics

random truncations were detected in two lines. One of these truncations was found to

occur in the 5’ upstream sequence between the KE and CE (∆KE6) (Figure 3-1A). These

mice retained tissue-specific expression of the downstream ETNK2 gene, but renin

expression was ubiquitous and unresponsive to ACE inhibition (Figure 3-1B). We

hypothesized that this was due to position effects of the insertion site because loss of the

KE on its own is not sufficient to explain these results. We therefore set out to

determine where the transgene inserted and if the renin gene had fallen under the

influence of that genes regulatory elements. Additionally, because of the retention of

ETNK2 expression, we questioned whether insulator elements resided around the renin

locus.

Methods

Generation of Kidney Enhancer-deficient PAC160 Transgenic Mice

Construction of the KE deletion mutant of PAC160 and of transgenic mice

containing PAC160 and PAC160∆KE6 were previously reported (147, 148). All mice

were fed with standard mouse chow (LM-485; Teklad Premier Laboratory Diets) and

water ad libitum. Care and use of mice met the standard set forth by the National

Institutes of Health and all procedures were approved by the University Animal Care and

Use Committee at the University of Iowa.

48

Transgene insertion mapping

Genomic DNA (gDNA) from a ∆KE6 mouse and a synthetically constructed

double stranded adapter were digested in separate reactions with BamHI, EcoRI, Nru I,

and Sal I for 4 hours. The sequences for all adapter and primer used are found in Table

3-1. Fragments were purified using Qiagen’s PCR purification kit and gDNA fragments

subsequently digested with Pac I for 4 hours. Separate overnight ligation reactions were

carried out with digested gDNA and the complimentary adapter. Aliquots of the ligations

were then used in PCR reactions with transgene and adapter specific primers. Reactions

were run on an agarose gel and an enriched fragment from the Sal I reaction was gel

purified and subjected to another round of PCR with nested primers. That reaction was

run on an agarose gel followed by gel purification of the enriched fragment. Purified

PCR product was then sent for sequencing with the nested primers (See Table 3-1) and

results BLATed against the mouse genome using the UCSC genome browser. Based on

the BLAT result, two forward primers were designed specific to the gene hypothesized to

contain the transgene insertion. Using those primers in combination with a transgene

specific reverse primer, PCR reactions were carried out using wild-type, PAC160, and

∆KE6 gDNA to confirm the site of transgene insertion. One ∆KE6 specific band was

then sequenced and BLATed to confirm the site of transgene insertion. We subsequently

used RT_PCR to assess the tissue expression of Zbtb20-human renin fusion transcripts

(See Table 3-1 for primer sequences). The RNase protection probe used to quantify

Zbtb20-renin fusion transcripts was derived from amplification of ∆KE6 RNA with the

primer set indicated and cloned into pCR4-TOPO2 (Invitrogen). Plasmids were

subsequently sequenced to confirm the identity of the probe.

RNA Isolation and RT-PCR

Tissues were homogenized in Tri-Reagent (Molecular Research Center,

Cincinnati, OH) and the RNA was isolated using chloroform phase separation and

49

isopropanol precipitation. Isolated total RNA was reverse transcribed and amplified

using the Superscript III One-step Reverse transcription (RT)-PCR with Platinum Taq kit

(Invitrogen) using the primers listed in Table 3-1.

In vitro Transcription Translation

Recombinant CTCF protein corresponding to the zinc finger domain

(hCTCFZnF11; ~45 kDA) was obtained using similar methods to those previously

described (150-152). Briefly, human cDNA encoding the full-length eleven zinc finger

domain of CTCF was amplified from Calu-6 cDNA and cloned into pQE-30 (Qiagen).

That cloned fragment was PCR amplified to generate a template for in vitro translation

using Platinum Taq DNA Polymerase High Fidelity (Invitrogen) and the primers listed in

Table 2-1. The presence of a single specific band was confirmed on a 1% agarose gel. A

5 µl aliquot of the PCR reaction was used as a template for in vitro transcription using

reticulocyte lysate TNT T7 Quick for PCR kit (Promega). Parallel 35S-Methionine

labeling reactions were carried out to confirm production of a specific protein product of

appropriate size. Labeled 35S protein was resolved on 10% SDS-PAGE gels.

EMSA

Electrophorectic mobility shift assays were performed as previously described

(150, 151, 153). Probes were constructed by annealing two single stranded oligos with

5’-GATC overhangs and the resultant double stranded probe was labeled with [α-32P]dATP. Probe sequences for the 5’ hypersensitive site 5 of the β-globin locus (HS5),

human renin intron (probe 1), and the intergenic sites between ethanolamine kinase 2

(ETNK2) and SRY-box 13 (SOX13) (probes 2 and 3) were obtained from the UCSC

genome browser. The HS5 probe has been previously described (153) and potential

CTCF binding sites were identified by loading a track accessible at http://bioinformatics-

renlab.ucsd.edu/rentrac/wiki/CTCF_Project from the lab of Bing Ren (151). Binding

reactions were carried out in 1X phosphate buffered saline solution with 5 mM MgCl2,

50

0.1 mM ZnSO4, 1 mM DTT, 0.1% Nonidet P-40, 10% glycerol, and 50 ng/µl of poly (dI-

dC) plus a 44-mer double stranded competitor (See Table 2-1). Each reaction contained

2 µl of programmed extract and 20-40 fmol of labeled probe. Competition reactions

contained 100-fold molar excess of cold probes. Each reaction was incubated at room

temperature for 30 minutes followed by separation of complexes on 5% polyacrylamide

gels in 0.5X TBE.

Chromatin Immunoprecipitation

Mouse liver tissue for ChIP assays was harvested from (∆KE6) transgenic and

non-transgenic mice. Tissues were minced on ice and suspended in PBS containing 1%

(v/v) formaldehyde at room temperature for 15 min. Reactions were stopped by the

addition of glycine (0.125 M, 5 min, room temperature), homogenized on ice, and then

rinsed with ice-cold PBS three times. The final washed pellet was resuspended in lysis

buffer with protease inhibitors (EZ-CHIP kit, Millipore) and sonicated on ice under the

following conditions (Amplitude 50%, time 15 seconds, cooling 15 seconds, 10-15 times

using a Sonic Dismembrator Model 500, Fisher Scientific). The size of the sonicate

chromatin was verified as between 400-700 bp by electrophoresis. The chromatin

immunoprecipitation was performed following the instructions provided by the

manufacturer using CTCF antibody (Upstate Biotechnology, Millipore, 07-729), rabbit

normal IgG as negative control and 2% input as a positive control. The primers used for

PCR are shown in Table 2-1. The mouse H19 gene was used as a positive control for

CTCF binding (154).

Results

To determine the site of transgene insertion we developed a cloning method that

would allow us to isolate the 5’ flanking gene (Figure 3-2A). Using information from

PCR assays we knew approximately were the truncation occurred. We designed an

51

adapter with four different restriction enzyme sites unique from a Pac I site close to the

transgene breakpoint. By performing two rounds of restriction digestion we were able to

ligate the adapter to fragmented genomic DNA (gDNA) from ∆KE6 mice. Adapter

ligated gDNA was then subjected to two rounds of amplification using adapter and

transgene specific primer resulted in isolation of three fragments (Figure 3-2B).

Sequencing of one of those and aligning those fragments to the mouse and human

genomes revealed that the transgene had inserted between exons 1 and 2 of the Zbtb20

gene on chromosome 16. We were also able to map the precise location of transgene

truncation to 10,440 bp upstream of the renin transcription start site. The alignment also

indicated that the direction of hREN and ETNK2 were the same as Zbtb20. Validation of

the insertion site was done using primers near the insertion site in Zbtb20 and near the

breakpoint in the transgene. Two separate primer pairs detected specific bands only in

genomic DNA of ∆KE6 mice (Figure 3-2C). Sequencing and alignment of these

fragments corroborated our previous results that insertion of the ∆KE6 transgene was in

Zbtb20.

We next sought to determine if the ubiquitous expression of renin was due to

initiation from the Zbtb20 promoter. Using a primer in exon 1 of Zbtb20 and exon 5 of

human renin, we performed RT-PCR on RNA from various tissues of ∆KE mice. We

detected the presence of two predominant transcripts that were ubiquitously expressed in

transgenic mice but absent in non-transgenic or wild-type controls (Figure 3-3). Gel

purification of the fragments from the heart sample, cloning, and sequencing revealed

that in fact there were three transcripts initiated in exon 1 of Zbtb20 (Figure 3-4). This

suggested that renin expression had fallen under the control of the Zbtb20 promoter. Of

note is fusion transcript number 1. Exon 1 from Zbtb20 has formed a fusion transcript

with exon 1b of the human gene plus an additional upstream exon (Figure 3-4). This

would suggest that expression of this exon has been deleted or also fallen under control

of the Zbtb20 promoter. In lieu of determining the expression pattern of Zbtb20 in wild-

52

type mice, a search of gene expression profiles from the Gene Expression Omnibus

database was performed. Every data set for brain, heart, lung, skeletal muscle, and testes

was positive for Zbtb20. This indicates that the ubiquitous pattern of Zbtb20 fusion

transcript expression mirrors that of the normal locus. To determine if transcription

initiated at the renin promoter was still intact, we performed an RNase protection assay to

quantify transcription initiation at the Zbtb20 promoter and the renin promoter. Using a

probe corresponding to the major band identified in RT-PCR, we determined transcript

levels in several tissues. Two protected transcripts were identified in all of the tissues

tested but not non-transgenic mice (Figure 3-5). The upper band is entirely Zbtb20-

hREN fusion mRNA and was identified only in the tissues of transgenic mice. There was

a weak band detected in wild-type mouse RNA that is probably the result of incomplete

digestion, but remains to be identified. The second band represents only human renin

sequence and was detected in all of the tissues tested. This represents primarily

transcripts controlled by the renin promoter. Therefore, transcription of renin is not only

being initiated at the Zbtb20 promoter, but its regulatory elements are influencing the

renin promoter.

Despite the ubiquitous pattern of renin expression, ETNK2 expression maintained

its localization to the kidney, liver, and testes. The deletion of sequences upstream of

renin, its dysregulation, and preservation of ETNK2 expression were indicative of the

presence of insulators around the renin locus. A genome-wide binding ChIP study from

Bing Ren’s group identified six CTCF binding sites in and around the human renin locus

(Figure 3-6A) (151). Since that analysis, the genome-wide ChIP identification of those

binding sites has been replicated in different cell types by several other groups including

those part of the Encyclopedia of DNA elements (ENCODE) project (155, 156). These

sites have also been identified in genome wide screens in the mouse indicating that CTCF

binding at these sites is conserved across species (157-159). These data strongly support

the existence of insulators in and around the renin locus. To validate these binding sites

53

we used 32P-lableled probes corresponding to sites identified in the renin intron (site 1)

and downstream of ETNK2 (sites 2 and 3) in EMSAs (Figure 3-6A). When incubated

with in vitro translated CTCF-zinc finger domain protein, all of the sites identified

previously provided a gel shift (Figure 3-6C). This gel shift was effectively competed

away by excess cold probe (S) that was not observed with mutant probe (M).

Furthermore, HS5 and another control probe, BR8, resulted in gel shifts (Figure 3-6D).

Another potential binding site identified by scanning the region between the renin and

ETNK2 genes (RE) for the CTCF motif identified by Kim et al., did not show binding.

Interestingly, a site identified by Kagey et al. just upstream of the mouse enhancer

(mR5’) showed weak binding (Figure 3-6D). This site does not show binding

conservation in any studies to date in the human and may represent a species-specific

event.

The binding sites around the renin locus clearly have the potential to bind CTCF.

We performed ChIP on chromatin from the livers of ∆KE6 mice and non-transgenic

controls. The three sites that showed CTCF binding and the proposed site between renin

and ETNK2 were amplified from CTCF antibody (αC) precipitated chromatin. The DNA

methylation free domain of the mouse H19/Igrf2 known to bind CTCF was included as a

positive control (160). Primers designed for site 1, site 2, RE, and mH19 all provided

positive signals from input (I) chromatin from transgenic animals (Figure 3-7C). Input

signals were not detected in non-transgenic liver chromatin for site 1, site 2, and RE

indicating that the assays designed were specific for the transgene. All of the regions

tested in this assay showed clear enrichment for CTCF immunoprecipation whereas the

IgG (-) had no signal from transgenic chromatin. The RE site showed weak binding as

indicated by its requirement of 40 cycles of PCR for detection versus the 30 cycles used

for other sites. This is consistent with the EMSA data where no shift complex was

observed despite sequence conservation with a consensus CTCF binding motif. The

sequence most likely dissociated during electorphoresis because of CTCF’s apparent low

54

affinity for the RE. However, CTCF binding to chromatin at the RE site was detected

and suggests that it could be functional. To quantify the binding of CTCF to each site

we performed the same immunoprecipitations and probed for the same sites with the

addition of a negative control (CD3). Site1, site 2, and mH19 had robust ChIP signals

(black bars) compared to the IgG control (grey bars) after correcting for differences in the

input signal. When comparing ChIP signals of the different sites to the negative control,

site 1 in the first intron of renin and site 2 downstream of ETNK2 show a ~300-fold and

~450-fold enrichment over the negative control. The signals for the mH19 and RE sites

were 45-fold and 21-fold enriched respectively. These data taken together indicate that

CTCF binds directly to sequences in the intron of renin, the intergenic regions between

renin and ETNK2, and the intergenic region between ETNK2 and Sox13.

Discussion

The regulation of human renin gene expression in the past has been difficult to

study due to the lack of a suitable cell line. Those problems are compounded by the large

5’ upstream sequence flanking the renin gene making it difficult to tease out important

cis-regulatory elements like the enhancer that reside several kilobases away from the

promoter. Those elements are typically taken out of their normal context so they are

amenable for transient transfection promoter bashing experiments. These have been

useful in identifying transcription factors that can regulate renin expression. Their scope,

however, is limited to viewing the regulation of transcription in a linear context where

there is a regulatory element such as an enhancer and it activates transcription of the

nearest promoter. With the advent of such techniques as chromosome capture

conformation, we have a better appreciation for the three dimensional regulation of

transcription through the interactions of regulatory elements hundreds of kilobases away

interacting with each other to affect gene expression. For that reason, our lab is utilizing

large genomic constructs like the PAC160 transgene used in this study. Because of the

55

extensive sequence flanking the gene renin there is a greater chance that all of the

elements for regulation are present. Here I have described the characterization of a

transgenic mouse line carrying a random truncation in the PAC160 construct that led to

ubiquitous renin expression. It allowed for the unbiased identification of CTCF binding

sites surrounding the renin gene that may be important for insulator enhancer-blocking

activity that maintains tissue- and cell- specific gene expression.

Previous hREN transgenic mice with short upstream sequences have failed to

faithfully recapitulate the regulation of renin in response to physiological regulators.

Surprisingly, all of these mice maintain their juxtaglomerular (JG) cell-specific

expression (145). If renin expression was not ubiquitous, then the responses to activating

or repressing inputs were dysregulated. The one commonality among all of the hREN

transgenics is the retention of JG specificity and the presence of the CTCF binding site in

the intron. Of particular interest is the 0.14 kb hREN transgenic mouse from our lab

(144). Despite having only 149 bp of the 5’ upstream sequence, they maintain cell-

specific expression within the kidney. This would suggest that there may be elements

within the structural gene itself that regulate cell-specific expression. The first intron of

renin might serve such a role, but appears to be insufficient on its own to regulate tissue-

specific expression. Previous studies have identified potential negative regulatory

elements within the intron (161). Others have observed a downregulation in reporter

gene assays when the human intron is included in constructs (162, 163). Our

identification of a CTCF binding site within the intron may account for that negative

regulation. In fact, CTCF was originally identified as a negative regulator of the c-myc

gene by using reporter gene assays (164, 165). Recent work has shown that CTCF within

an intron of the PUMA gene acts as a repressor of p53 transcriptional activation. It

appears to function by impeding pol II movement through the CTCF binding site that

results in constitutive production of a long RNA transcript. Of note is the co-localization

of the cohesin complex with CTCF in these intragenic sites. Cohesin has been found to

56

co-localize with CTCF at insulators and mediate enhancer blocking activity through the

formation of intrachromosomal loops (166-168). If the intron site is involved in cell-

specificity, it is most likely not due to an interaction upstream of the renin gene as

sequences past 146 bp from the transcription start site appear to be dispensable.

Genome-wide ChIP in mouse ES cells shows a signal for components of the cohesin

complex that overlaps the CTCF binding in the renin intron (159). Whether or not loop

formation is playing a role in blocking pol II remains to be seen. Also, searching

ENCODE data using the UCSC genome browser indicates the presence of a DNase I

hypesensitive site. What makes this noteworthy is its existence in cells not thought to

express renin at high levels if at all. One might hypothesize that this is a constitutive

DNaseI hypersensitive site that allows for CTCF binding in multiple cell types to repress

renin expression by blocking the progression of pol II. Is CTCF the cell-specific

determining factor in the kidney? It may be since cell-specific expression was

maintained in the ∆KE6 mice and other hREN transgenics (169). We have the tools

available with BAC transgenesis to delete this sequence and test the effects in transgenic

mice within the native context of the gene.

The preservation of ETNK2 expression in the face of renin dysregulation and the

existence of a CTCF binding site strongly support the presence of an enhancer between

hREN and ETNK2. The weak binding interaction between the RE and CTCF probably

explains why genome-wide ChIP assays have failed to identify this site. Our ChIP-qPCR

method is more sensitive than genome tiling arrays or high throughput sequencing and

was able to detect binding in the RE region. The binding of this site by CTCF and the

downstream intergenic site between ETNK2 and SOX13, or the intragenic hREN site,

might allow for formation of a chromatin loop like those formed in β-globin gene

activation (170, 171). This loop formation may protect ETNK2 from the influences of

regulatory elements in neighboring genes. In the case of ∆KE6 mice, loop formation

57

would shield ETNK2 from the influences of the Zbtb20 gene that result in the

dysregulation of renin.

Our data from ∆KE6 mice suggest that the loss of sequences upstream of the

hREN transgene render it susceptible to positional influences. Whether it is due to the

loss of an insulator at the CTCF binding site in the GOLT1A gene identified by others

remains to be determined. We did not validate CTCF binding at this site in full length

PAC160 mice. However, it is attractive to speculate that an insulator at that site is

responsible for protecting the tissue-specific expression of renin. It is entirely possible

that there is an element lost in ∆KE6 mice that normally represses renin in non-renin

expressing tissues, but our RT-PCR and RPA data argue against this. Furthermore, the

lack of a response to ACE inhibition suggests that Zbtb20 regulatory elements are

exerting the predominant role in promoter activity given that 0.14 kb hREN transgenic

mice are able to respond appropriately. It has now been observed multiple times that loss

of a large portion 5’ upstream sequence of the renin gene vulnerable to position effects.

It now needs to be determine whether the CTCF binding sites within and around the renin

gene are within functional enhancers to serve to protect appropriate renin transcription

activity. Utilization of our PAC constructs to delete these elements in the normal

genomic context will serve as a valuable tool.

58

Table 3-1. Primers and probes for determining gene expression, transgene insertion mapping, EMSA, and ChIP analysis.

59

Figure 3-1. Stucture of the PAC160 and ∆KE6 transgenes and renin expression.

A) Genes and hREN regulatory sequences contain within the PAC160 transgene. Displayed below is the ∆KE6 transgene showing the breakpoint 10.44 kb upstream of the renin transcription start site. B) RNase protection assays measuring the expression of GOLT1A, ETNK2, and hREN expression in PAC160 and ∆KE6 mice at baseline. Below is the hREN and mRen expression after captopril treatment.

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Figure 3-2. Mapping the ∆KE6 insertion site in the mouse genome.

A) Schematic representation of the general strategy to map transgene insertion within the mouse genome. B) (left) Agarose gel of ligation fragments from the first round of PCR following genomic DNA digestion with Bam HI (B), Eco RI (E), Nru I (N), or Sal I (S) and ligation to the equivalently digested adapter. The last lane is BamHI digested

tion fragments following PCR cleanup or gel purification in the case of Sal I (P) fragment from the left gel. C) Confirmation of the transgene insertion site using primer sets in the Zbtb20 locus and ∆KE6 transgene. N= non-transgenic W= PAC160 ∆6= ∆KE6

genomic DNA minus the adapter. (right) Nested PCR amplified liga

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Figure 3-3. Detection of fusion transcripts in ∆KE6 mice.

RT-PCR using primers in exon 1 of the Zbtb20 gene and exon 5 of hREN. NT=transgenic WT2 and WT1= PAC160 transgenic

non-

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Figure 3-4. Genome structure and transcripts of ∆KE6 mice.

Red lines indicate the truncation and insertion of ∆KE6 or Zbtb20 respectively. This displays the different Zbtb20 transcripts and Zbtb20-hREN fusion transcripts as determine by cloning and sequencing the RT-PCR products from ∆KE6 mRNA. This figure also shows the alignment of hREN exon 1b transcript to that of fusion transcript 1.

63

Figure 3-5. RNase protection assay identifying fusion versus hREN transcripts.

A probe was generated from the primary band from RT-PCR. The 246 bp band represents Zbtb20-hREN fusion transcripts and the 197 band detects only renin transcript. NT= non-transgenic WT2= PAC160 transgenic ∆KE2= full length enhancer knock out

64

Figure 3-6. EMSA analysis of identified CTCF binding sites.

A) Diagram of intact PAC160 transgene and the CTCF binding sites identified by Kim et al. B) SDS-PAGE gel of in vitro translated CTCF-zinc finger domain. 35S-Methionine labeled programmed (+) or unprogrammed (-) extract. C) Gel shift analysis of programmed extracts incubated with 32P-labeled double-stranded oligos for the identified binding sites 1, 2, 3, and RE. Known binding sites of CTCF HS5 and BR8 were included as positive controls. The mR5’ probe represents a conserved CTCF binding motif upstream of the enhancer. Excess cold probe (S) or mutant (M) was used for competitions. Labeled mutant probes do not bind CTCF (M2 and 3).

65

Figure 3-7. ChIP analysis for the binding of chromatin around the ∆KE6 transgene.

A) Location of CTCF binding sites within the ∆KE6 transgene B) ClustalW alignment and box shading for the CTCF consensus motif from Kim et al. and identified binding sites within PAC160. Shading was done using a server located at http://www.ch.embnet.org/software/BOX_form.html running Boxshade 3.21. Black represents a base identical to the consensus sequence and gray indicates a similar base. C) ChIP for CTCF binding sites for the mH19 ICR region and sites within the transgene

for quantification of enrichment. Left, ChIP signal after subtraction of the input chromatin signal. Right, Fold enrichment relative to the CD3 negative control locus

site 1, site 2, and RE. D) ChIP performed as in C but followed by qPCR

66

CHAPTER 4

DICER MAINTAINS THE RENIN CELL PHENOTYPE

Introduction

The renin-angiotensin system (RAS) has been established to be important in the

regulation of kidney development. Knockout of RAS components in mice results in

lethality typically between birth and weaning (172-176). Humans with mutations that

inactivate these genes typically die in utero (177). The RAS components are thought to

modulate cell proliferation and differentiation during development or prevent ischemia in

the kidneys. Renin is involved in the rate-limiting step of the RAS and initiates the

cascade of events that leads to angiotensin II production that is important for blood

pressure, electrolyte/fluid, and metabolic homeostasis. Renin knockout mice develop

severe hydronephrosis, cortical atrophy, shrinkage of tubules, interstitial fibrosis,

glomerular sclerosis, and hypertrophy of the kidney vasculature due to increases in cell

numbers (174, 177). These are common features of RAS knockout animals (174, 178-

180). Notably, mice in which the renin cell itself is ablated do not display this

phenotype. This suggests that the renin cell itself or something else that it secretes is

responsible for the hypertrophy. Therefore, the cells surrounding the vessels in RAS

knockout mice are probably cells that were destined to become renin expressing cells.

Small RNAs, particularly microRNAs (miRNA), have become a major area of

research because of their diverse functions. Notably, miRNAs are involved in a variety

of physiological pathways and diseases by regulating gene expression. They are first

transcribed as 100-1000 nt long RNAs that are trimmed by Drosha/DGCR8 in the

nucleus and then transported to the cytoplasm by exportin-5. The enzyme Dicer then

processes them for loading into the RNA induced silencing complex (RISC) which the

miRNA guides to target mRNAs to carry out gene repression. They do this primarily

through repression of translation, but can also cause RNA degradation.

67

The development of several tissues and organs are regulated by miRNAs (181-

185). The knockout of Dicer using specifically expressed cre recombinase expressing

lines and the Dicer-flox mouse expressing lines has been utilized by others to determine

if miRNAs are involved in development or disease because its deletion results in the

absence of miRNA expression in the targeted tissue or cell-type. The deletion of Dicer

has been performed in cells of the nephron lineage epithelium and the collecting duct

epithelium (186-188). It results in the termination of nephrogenesis or severe kidney

abnormalities. When specifically deleted in podocytes, loss of dicer results in the

progression of glomerular disease (186, 187). Given their role in the development of

other tissues and cell-types, we were interested in determining what role microRNAs

(miRNA) play in the maintenance of the renin expressing cell.

Methods

Generation of Dicer KO mice

Mice expressing Cre recombinase from the endogenous renin locus Ren1dcre/+

were bred to mice homozygous for the Dicer floxed allele (Dicerflox/flox). A second cross

of Ren1dcre/+; Dicerflox/+ offspring were bred to Dicerflox/flox to produce Ren1dcre/+;

Dicerflox/flox knockout mice. Alternatively, because Cre may not efficiently delete both

alleles, heterozygous knockout mice Dicer∆/flox were bred to Ren1dcre/+ so that Cre only

had to delete one allele (189, 190). Control mice were Dicerflox/+; Ren1dcre/+ to ensure

comparisons were done in mice with the same number of renin gene alleles.

All mice were fed with standard mouse chow (LM-485; Teklad Premier

Laboratory Diets) and water ad libitum. Care and use of mice met the standards set forth

by the National Institutes of Health and all procedures were approved by the University

of Iowa Animal Care and Use Committee at the University of Iowa.

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Histological analysis and immunostaining

Mice were CO2 asphyxiated, kidneys extracted and placed in Bouin's fixative

overnight. Paraffin sections were stained with hematoxylin, with PAS and Masson's

trichrome as previously described (191). Immunostaining was performed on 5 µm thick

paraffin sections for renin (1:500 dilution of rabbit anti mouse renin polyclonal antibody)

and α-SMA (1:10,000 dilution of a monoclonal anti- α-SMA-specific antibody isotype

Ig2a; Sigma, St. Louis, MO) as previously described (190).

RNA extraction and quantitative RT-PCR (qRT-PCR) analysis

Kidneys were homogenized in Trizol (Invitrogen) and phase separation was

performed using chloroform. RNA was extracted from the aqueous phase using

Purelink™ RNA Mini Kit (Invitrogen) with on-column DNase treatment. First strand

cDNA was generated from 1 µg of total RNA using Superscript III (Invitrogen) in a total

reaction volume of 20 µl. The first strand reaction was diluted 1:45 and 9 µl of the

dilution was used for quantitative real-time PCR. Taqman® assays were run using primer-

probe sets and master mix from Applied Biosystems- Ren1 (Assay ID-

Mm02342888_gH), Ren2 (Assay ID- Mm00651435_mH), and β-Actin (Part #-

4352933E). The specificity of Ren1 and Ren2 probes was tested using cloned cDNAs for

each gene. Relative expression was determined using the 2-∆∆Ct method.

Tail cuff and Telemetry BP measurements.

Mice were trained for tail cuff measurements for 7 days followed by daily

recording for an additional 7 days using a BP-2000 apparatus (Visitech System Apex,

Inc.). For recording each day, there were 10 unrecorded cuff inflations and 30 recorded

inflations. An average of at least 20 successful recordings were used for data analysis.

Radiotelemetry was performed as previously described (176).

69

Results

Using allele-specific qRT-PCR for Ren1 and Ren2, we determined the levels of

expression in control and JG cell selective KO animals (Figure 4-1A). Knockout mice

show a significant reduction in the levels of renin mRNA. Expression of Ren1 was

reduced to 20% of the control and Ren2 to approximately 10%. To determine whether or

not expression of the renin genes was still inducible, we treated mice with the ACE

inhibitor Captopril. Control mice responded robustly with a 19-fold induction whereas

the KO mice had a modest increase of about three-fold (Figure 4-1B). Immunostaining

for renin shows a dramatic reduction in the number of renin positive JG cells (Figure 4-

1C). When quantified, the percentage of JG cells expressing renin is 33 ± 4.37%

compared to the knockout mice where just 1.43 ± 0.35% were positive. The decrease in

the number of renin expressing cells in the kidney most likely causes the decrease in

renin expression. However, renin cells are capable of upregulating renin expression in

KO mice since there was still a response to angiotensin converting enzyme inhibition,

albeit much lower than wild-type. Whether or not it is due to an increase in expression

from that 1% of cells with preserved renin expression or a recruitment of non-renin

expressing cells was not determined. Previous results from angiotensinogen deficient

mice suggest that it would be a recruitment (179).

Not surprisingly, there was a decrease in the blood pressure of KO mice when

compared to controls. Tail cuff measurements of Dicer∆/flox; Ren1dcre/+ mice revealed a 15

mmHg decrease in their blood pressure versus controls (Figure 4-2A). Telemetry blood

pressure measurements in Dicerflox/flox;Ren1dcre/+ indicated a 9 mmHg difference (Figure

4-2B). The difference may be due to the severity of the phenotype between the different

KO genotypes, but the tail cuff method may have overestimated the blood pressure. Only

a sample size of three for each mouse was measured by tail cuff, whereas six for each

group were used in telemetry measurements. Given the small sample size it is difficult to

draw conclusions about the severity of the phenotype. There were also difficulties in the

70

radiotelemetry studies as implantation of the telemeter in Dicer∆/flox; Ren1dcre/+ mice

caused lethality.

Kidneys from KO mice displayed several abnormal phenotypes. Probably the

most obvious is the “bumpy” appearance of KO mouse kidneys. The bumps are formed

because of cortical indentations corresponding to stripes of fibrosis evidenced by

Mason’s trichrome staining (Figure 4-3A). Periodic acid-Schiff staining revealed that the

border between the cortex and medulla was poorly defined (Figure 4-3B). In addition,

the medulla appears to be atrophied. Immunostaining for α-smooth muscle actin (α-

SMA) indicates a marked increase in the expression of α-SMA (Figure 4-3C). However,

those areas of increased expression do not contain an increase in the number of cells, but

an increase in fibrosis around the vessels. Interestingly, this appears to be a common

phenotype with renin cell-specific knockout CBP/p300 mice where renin cells fail to

develop (191).

Discussion

The deletion of Dicer in renin expressing cells of the kidney results in several

kidney abnormalities. Most notable is the dramatic loss in the number of renin

expressing JG cells. This indicates an important role for Dicer-dependent miRNAs in

maintenance of renin expressing cells during development. The loss of JG cells was

associated with a reduction in the blood pressure. This reduction would normally induce

expression of renin, but it appears insufficient in Dicer knockout mice. However, renin

expression is stimulated by the treatment of mice with an ACE inhibitor. This activation

indicates that some cells may have escaped the deletion of Dicer. This is a possibility

given that this analysis was done in Ren1dcre/+; Dicerflox/flox knockout mice and requires the

deletion of two alleles versus the one in the Ren1dcre/+; Dicer∆/flox knockout line.

Recombinase deletion of two alleles may not have been 100% efficient. Indeed the blood

pressure phenotype appeared to be less severe. However, renin mRNA expression does

71

not appear to be different when looking at the individual values in qPCR analysis after

normalization to β-actin. Another scenario may be that given the structural abnormalities

of the kidney, particularly the perivascular fibroplasia, the baroreceptor function that

normally responds to changes in perfusion pressure is altered. Whether the reduction in

renin expression is secondary to the structural changes of the kidney is a critical question

to answer. It does appear that some of the abnormalities are due to a reduction in renin

expression as indicated by the medullary atrophy seen in other RAS knockouts.

Measurement of renin expression early during kidney development will be important for

addressing this question.

The kidneys of Dicer knockout mice display some unique features not observed

when components of the RAS are deleted. For instance, the striped pattern of fibrosis is

not observed in RAS knockout mice. Those mice display a more diffuse pattern of

interstitial fibrosis. This fibrosis in Dicer knockout mice may be more likely a result of

miRNA loss rather than the loss of renin expression. Recently, microRNA 200a (miR-

200a) was found to regulate the expression of transforming growth factor beta (TGF-β)

through a direct interaction with the 3’ UTR of the mRNA (192). The overexpression of

miR-200a in proximal tubule epithelial cells prevented the expression of pro-fibrotic

genes and TGF-β induced epithelial to mesenchymal transition. Moreover, the

expression of miR-200a was reduced in mouse models of diabetic nephropathy in which

progressive fibrosis observed. A model in which the loss of miR-200a in Dicer knockout

mice leads to increases in the pro-fibrotic TGF-β levels is attractive. In fact, a loss of

renin and therefore angiotensin II would be expected to result in a loss of TGF-β (193). It

will be of interest to examine the levels of TGF-β in the kidneys of Dicer knockouts.

Even though there appears to be renin cell-independent effects on the progression

of the kidney phenotype in Dicer knockout mice, the renin cell is involved. The

phenotype of the smooth muscle cells would support the previously established role for

renin cells. Like mice where renin cells are ablated, the vasculature of the kidney does

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not display hypertrophy like that seen in RAS knockouts. The cells surrounding the

vessels contributing to hypertrophy are most likely renin-expressing cells like those seen

in ACE knockout mice (178). Thus, the reduction in renin expressing cells might lead to

decreases in the release of another factor besides renin that stimulates the proliferation of

smooth muscle cells.

The data presented here indicate that Dicer-dependent miRNAs are important in

the maintenance of the renin cell phenotype. There appear to be both renin cell-

dependent and -independent mechanisms controlling the abnormal kidney phenotype. It

will be interesting to determine what miRNAs are differentially expressed in renin

expressing cells and those of the renin lineage. This may identify genes that are

important for the transition of a renin expressing cell into a smooth muscle cell.

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Figure 4-1. Expression of Ren1 and Ren2 in mouse kidneys of control and knockout mice.

A) Expression of Ren1 and Ren2 in the kidneys of control and knockout mice as determined by qPCR (N=6 per group; * = p<0.05; t-test). B) Induction of Ren1 expression in response to ACE inhibition in control and knockout mice. Y-axis is in log scale. (N=3 for each control group; N=2 for each knockout group C) Kidney immunostaining for renin protein. Bar= 200 µm

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Figure 4-2. Blood pressure measurements in control and Dicer knockout mice.

A) Tail cuff measurement of systolic blood pressure (SBP) (N=3 for each group, * = p<0.05). B) Radiotelemeter measurement of arterial pressure plotted as the mean arterial pressure (MAP) (N=6 for KO; N=7 for control; * = p<0.05; t-test).

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Figure 4-3. Histology of control and Dicer knockout mice.

A) Mason’s trichrome staining of control (left) and Dicer knockout (right) mice for assessment of collagen deposition (blue) and fibrosis. B) Periodic acid Schiff staining for membrane integrity. Arrows indicate cortical indentations. C) α-smooth muscle actin staining (brown) for marking vascular smooth muscle cells.

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CHAPTER 5

GENERAL DISCUSSION

Renin Expression and Maintenance of the Renin Cell Phenotype

Summary of Results

My studies have focused on three different aspects of the regulation of renin.

First, I have examined the role of the orphan nuclear receptors Nr2f2, Nr2f6, and Nr4a1

in regulating renin expression. Both Nr2f2 and Nr2f6 are negative regulators of the renin

promoter and can bind to the hormone response element of the enhancer to mediate that

effect. However, Nr2f2 does not regulate baseline expression of the endogenous gene.

The orphan Nr4a1 acts as a positive regulator, but it does not do so through direct binding

to the HRE. Future experiments will be aimed at determining what signals can activate

Nr2f2 and its actions and how Nr4a1 might exert its function.

Second, I have identified CTCF binding sites in and around the renin locus. The

insertion of a truncated phage artificial chromosome (PAC) transgene in the mouse

Zbtb20 gene resulted in ubiquitous renin expression. However, ETNK2 maintained its

normal pattern of expression. A CTCF binding site between renin and ETNK2 probably

represents an insulator that shields ETNK2 from the influences of regulatory elements in

the Zbtb20 gene. Because of the loss of 5’ upstream sequences in the PAC transgene,

human renin expression downstream of the truncation was driven by regulatory elements

of the Zbtb20 gene. The upstream sequence likely responsible is a CTCF binding site

identified by other groups that was deleted in the broken transgene. Whether the CTCF

binding sites are acting as insulators and what impact they have on renin expression will

be the target of further experimentation.

Third, I have investigated the role of Dicer-dependent microRNAs (miRNAs) in

controlling the maintenance of the renin expressing cell phenotype. The deletion of Dicer

results in a dramatic reduction in the number of juxtaglomerular (JG) cells and severe

77

kidney abnormalities. Which miRNAs are important for the development of smooth

muscle cells that are derived from a renin cell precursor and which are needed for proper

development of renin expressing cells is the main question yet to be answered.

Future Directions

Although each one of the projects above has their own questions still to be

answered, we do not have knowledge of other regulatory elements and those that mediate

the responses to physiological cues. I will therefore focus on a series of experiments that

I believe will contribute generally to the understanding of renin expression regulation.

Each one of the aspects of renin expression studied above is probably impacted at the

level of chromatin. Histone modifications like acetylation and methylation control the

accessibility of DNA for transcription factor binding by affecting nucleosome occupancy.

They can also act as marks for the recruitment of transcription factors. What chromatin

modifications exist at regulatory regions during renin activation or repression remain

largely unexplored. Additionally, it is not known what other sites are accessible to

transcription factors that act as regulatory elements apart from the currently identified

enhancer and promoter. It is now clear that the enhancer functions as a baseline regulator

of renin expression but not an element that controls renin expression in response to

physiological input. The enhancer likely regulates the magnitude of the inhibition or

activation but is not necessary for an intact physiological response. This is evidenced by

the observation that deletion of the enhancer retains physiological responsiveness.

Therefore, it will be important to define the regulatory elements and transcription factors

acting as the primary regulatory elements responding to signals that regulate renin

expression. Identification of those regulatory elements is critical for understanding how

renin expression is regulated by changes in blood pressure and electrolytes.

In order to identify other regulatory elements it will first require determination of

the regions that are accessible to transcription factor binding. Formaldehyde assisted

78

isolation of regulatory elements (FAIRE) is more amenable for probing the large 5’

upstream sequence of renin over traditional methods like DNase hypersensitivity for

finding accessible regions (194). It exploits differences in the crosslinkability of histones

versus other proteins to DNA. After formaldehyde crosslinking of cell chromatin,

nucleosome free regions of active gene promoters preferentially segregate into the

aqueous phase upon phenol-chloroform extraction. Sites representing nucleosome free

regions can be probed for enrichment using PCR. This is much more amenable for

screening a large upstream region like that of renin rather than using Southern blots to

identify DNase hypersensitive sites. Using this approach combined with sequence

conservation information will allow for the identification of new regulatory elements that

may be the regulators of renin expression in response to physiological cues.

Identification of nucleosome free regions as a first step would be canonical to the

characterization of the β-globin gene (195-198). Regulatory regions in that gene were

initially identified using DNase hypersensitivity assays. That analysis localized several

nucleosome free regions that have now been found to be important in cell-specific

regulation of the β-globin locus. At least two of these regions are CTCF insulator

binding sites that exhibit enhancer blocking activity. Another is an insulator with barrier

activity that does not require CTCF. Identifying accessible regions in and around the

renin locus will allow for the prioritization of regions for future studies. Of course the

potential CTCF binding sites would be important sites to test initially. Additionally,

primer sets spanning the renin intron and 5’ upstream sequence of renin could be

designed to explore those regions. This analysis would not only be useful in As4.1 cells

that express renin at a very high level, but also non-renin expressing cells that can be

induced to express renin. The sites accessible in both As4.1 cells and those induced to

express renin, but not in non-renin expressing cells, would be attractive candidates for

regulatory elements important for recruitment and maintenance of renin expressing cells.

This may be of particular interest in the case of renin expressing cell recruitment since

79

histone acetylation appears to play an important role (199). Those acetylated regions are

most likely those that become accessible to transcription factor binding and regulate

induction of renin expression.

Once accessible regions are identified, the transcription factors that bind them

must be identified. This could be accomplished through the use of the DAPA assay we

have developed combined with stable isotope labeling of amino acids in cell culture

(SILAC). This method utilizes the differential labeling of two cell populations. One

population is labeled with a stable isotope tagged amino acid and the other is not.

Nuclear extracts from each cell population are incubated with biotin labeled double

stranded DNA probe corresponding to the site of interest or a mutant probe. Proteins

binding to each probe are precipitated with streptavidin beads, mixed together, and run on

a mass spectrometer. The stable isotope labeling provides enough of a shift in peptide

peaks that those from wild-type and mutant probes can be identified. If a peak for the

wild-type shows greater abundance over the mutant probe peak it indicates a binding

protein for that sequence. That peptide can then be searched for in protein databases for

identification. This approach has been shown to be very successful at identifying

proteins that show differential DNA binding to sequences differing by a single nucleotide

(200). Not only can it be applied to new regulatory elements but it can also be used for

the HRE and proximal promoter element. Additionally, extracts from cell populations

treated with different stimuli or inhibitors could be tested for transcription factors that

show enrichment for binding under different conditions.

Once potential regulatory elements and the proteins that bind them have been

identified, their functional relevance must be assessed. In order to do this we can utilize

the BAC/PAC recombineering methods already developed in our lab to delete these

sequences. By creating transgenic mice from these constructs, the role of the deleted

elements can be determined by assessing the response of renin to physiological cues.

Alternatively, stable transfections using the modified PACs could be made. Those cells

80

could then be subjected to cAMP induction or stimulation of increases in intracellular

Ca2+. These two stimuli are known to have an important role in the physiological

regulation of renin expression. However, a suitable cell line may be difficult to choose

given the lack of cAMP response in As4.1 cells. Of course the first sequences that should

be deleted using BAC recombineering and transgenic production are those CTCF binding

sites that we have identified.

Utilization of the methods above would allow for us to move beyond analysis of

just the enhancer and promoter. Although important for the regulation of renin

expression they do not represent the only elements involved. In order to move forward in

determining how renin is regulated, those sites that can act as regulatory elements must

be identified. Additionally, sequences that define the renin locus must be identified to

narrow the region of interest. It is most likely to consist of the region that spans from the

CTCF binding sites in GOLT1A to the renin-ETNK2 intergenic site (Figure 2-6). Once

the borders of the renin locus are identified we can localize the elements within them that

regulate renin expression. The difficulty is in identifying the sites responsible for the

induction or repression of renin expression rather than just the baseline expression. It is

therefore attractive to create transgenic mice in which cells expressing or capable of

expressing renin are in their native context making contacts with cells that may be

important for the transmission of regulatory signals. We have developed the tools

necessary to identify new regulatory elements and transcription factors, now we must

apply them to advance the field further.

81

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Regulation of renin gene expression by CTCF, Nr2f2, Nr2f6, Nr4a1