[methods in enzymology] g protein coupled receptors - modeling, activation, interactions and virtual...

CHAPTER THIRTEEN

Biasing the Parathyroid HormoneReceptor: Relating In Vitro LigandEfficacy to In Vivo BiologicalActivityKathryn M. Appleton*,‡, Mi-Hye Lee*, Christian Alele*,Christine Alele*, Deirdre K. Luttrell†, Yuri K. Peterson‡,Thomas A. Morinelli†,}, Louis M. Luttrell*,},1*Division of Endocrinology, Diabetes & Medical Genetics, Department of Medicine, Medical University ofSouth Carolina, Charleston, South Carolina, USA†Division of Nephrology, Department of Medicine, Medical University of South Carolina, Charleston,South Carolina, USA‡Department of Pharmaceutical & Biomedical Sciences, College of Pharmacy, Medical University of SouthCarolina, Charleston, South Carolina, USA}Research Service, Ralph H. Johnson Veterans Affairs Medical Center, Charleston, South Carolina, USA1Corresponding author: e-mail address: [email protected]

Contents

1.

MetISShttp

Introduction

hods in Enzymology, Volume 522 # 2013 Elsevier Inc.N 0076-6879 All rights reserved.://dx.doi.org/10.1016/B978-0-12-407865-9.00013-3

230
2. Determining the Relative Activity of PTH1R Ligands 232
2.1
Defining reference and test ligands 234 2.2 Assaying hPTH1R-mediated cAMP production 236 2.3 Assaying PTH1R-mediated intracellular calcium influx 243 2.4 Assaying PTH1R-mediated ERK1/2 activation 247 2.5 Estimating PTH1R ligand bias 254
3.
Discussion 256 Acknowledgments 258 References 259
Abstract

Recent advances in our understanding of the pluridimensional nature of GPCR signalinghave provided new insights into how orthosteric ligands regulate receptors, and howthe phenomenon of functional selectivity or ligand “bias” might be exploited in phar-maceutical design. In contrast to the predictions of simple two-state models of GPCRfunction, where ligands affect all aspects of GPCR signaling proportionally, currentmodels assume that receptors exist in multiple “active” conformations that differ in theirability to couple to different downstream effectors, and that structurally distinct ligandscan bias signaling by preferentially stabilizing different active states. The type 1 parathy-roid hormone receptor (PTH1R) offers unique insight into both the opportunities and

229

http://dx.doi.org/10.1016/B978-0-12-407865-9.00013-3

230 Kathryn M. Appleton et al.

challenges of exploiting ligand bias in pharmaceutical design, not only because numer-ous “biased” PTH analogs have been described but also because many of them havebeen characterized for biological activity in vivo. The PTH1R has pleiotropic signal-ing capacity, coupling to Gs, Gq/11, and Gi/o family heterotrimeric G proteins, and bindingarrestins, which mediate receptor desensitization and arrestin-dependent signaling.Here, we compare the activity of six different PTH1R ligands in a common HEK293 cellbackground using three readouts of receptor activation, cAMP production, intracellularcalcium influx, and ERK1/2 activation, demonstrating the range of signal bias that can beexperimentally observed in a “typical” screening program. When the in vitro activity pro-files of these ligands are compared to their reported effects on bone mass in murinemodels, it is apparent that ligands activating cAMP production produce an anabolicresponse that does not correlate with the ability to also elicit calcium signaling. Para-doxically, one ligand that exhibits inverse agonism for cAMP production andarrestin-dependent ERK1/2 activation in vitro, (D-Trp12, Tyr34)-bPTH(7–34), reportedlyproduces an anabolic bone response in vivo despite an activity profile that is dramat-ically different from that of other active ligands. This underscores a major challenge fac-ing efforts to rationally design “biased” GPCR ligands for therapeutic application. While itis clearly plausible to identify functionally selective ligands, the ability to predict howbias will affect drug response in vivo, is often lacking, especially in complex disorders.

1. INTRODUCTION

Early efforts to model the action of drugs or hormones assumed that

individual receptors behave as binary switches, existing in equilibrium

between an “off” state, which is silent in the assay, and an “on” state, which

is capable of generating a measurable response. In such models, receptor

conformation is the minimal determinant of system response and ligands

act solely by changing the fraction of the receptor population in the on state

(Karlin, 1967; Thron, 1973). The efficacy of a ligand thus becomes a reflec-

tion of its ability to stabilize the on state and can be approximated by two

parameters: the maximal observed response (Emax) and potency (EC50),

the ligand concentration that produces a half-maximal response. In this con-

text, full agonists are ligands that preferentially bind and stabilize the on state,

producing the maximum system response at saturating ligand concentration;

partial agonists are ligands with less conformational selectivity, translating into

a submaximal system response at saturating concentration and potential

attenuation of full agonist activity; true neutral antagonists are ligands with

equal affinity for both the off and on conformations, producing no physio-

logical response but able to block the response to agonists; and inverse agonists

are ligands that preferentially bind the off state, which causes them to appear

231Biasing the Parathyroid Hormone Receptor

as antagonists in systems with low basal activity but with the added property

of reducing receptor-mediated constitutive activity in systems with high

basal tone.

Although readily determined experimentally, measurements of EC50 and

Emax are limited in that they are influenced by system factors external to the

ligand–receptor unit (Ehlert, 2000; Figueroa, Griffin, & Ehlert, 2009). In the

case of G protein-coupled receptors (GPCRs), differences in receptor reserve

and signal amplification can lead to apparent changes in ligand classification

when comparisons are made between different assays. New signaling

responses commonly emerge as the level of receptor expression increases,

permitting less efficiently coupled effectors to reach the detection threshold

of the assay (Zhu, Gilbert, Birnbaumer, & Birnbaumer, 1994). Similarly,

variation in the expression levels of G proteins, arrestins, and downstream

effectors can make ligand activity appear to change between cell types

(Nasman, Kukkonen, Ammoun, & Akerman, 2001). Even in the same cell

background, ligands may appear as full agonists when classified using sig-

nals that are highly amplified, for example, cAMP production, but as par-

tial agonists when assayed for less amplified responses, for example, arrestin

binding and signaling (Rajagopal et al., 2010).

Nonetheless, signal strength arguments cannot account for true reversal of

potency or efficacy, for example, when the rank order of potency for two

ligands acting on the same receptor is opposite in two different assays of cellular

response (Berg et al., 1998). In a two-state model, ligand binding can alter the

fraction of receptors in the on state, but cannot qualitatively change the nature

of that state. Thus, the classification of a ligand as an agonist, antagonist, or

inverse agonist must be independent of the assay used to detect receptor

activation, and the rank order of potency for a series of ligands cannot vary

when two or more assays are employed. Reversal of potency or efficacy

implies that different ligands are activating the same receptor in different ways,

meaning that theymust be generating different active receptor states (Kenakin,

1995). That this phenomenon has now been described for several GPCRs,

among them the serotonin 5-HT2c, pituitary adenylate cyclase-activating

polypeptide, dopamineD2, neurokininNK1,CB1 cannabinoid,b2 adrenergic,angiotensin AT1A, and PTH1Rs, suggests that most, if not all, GPCRs can

adopt multiple active conformations (Luttrell & Kenakin, 2011).

It is nowapparent thatGPCRsignaling is “pluridimensional” (Galandrin&

Bouvier, 2006), meaning that receptors signal by coupling to multiple

G protein and non-G protein effectors. If different active conformations


couple the receptor to these downstream effectors with different efficiency,

then the cellular response will be dictated by the distribution of receptors

across the range of achievable active and inactive states. Since there is no

a priori reason that the active conformation(s) favored by one ligand should

be identical either to the spontaneously formedactive state or to those preferred

bya structurallydistinct ligand, thepotential exists for the ligand tobias signaling

in favorof someeffectors at theexpenseofothers.Thus, it is the ligand–receptor

complex, not the receptor alone, that specifies the active state, along with any

other smallmolecule, protein–protein, or lipid–protein interaction that alloste-

rically constrains the conformations available to the receptor (Kenakin &

Miller, 2010). In contrast to a two-state model, wherein agonists and antago-

nists merely control the quantity of receptor activity, the potential of biased

agonism lies in its ability to qualitatively change signaling.

Here, we employ the PTH1R to illustrate some of the issues arising from

pluridimensional efficacy and the challenge of adequately describing ligand

bias. Using a selected panel of PTH analogs and a multiplexed set of cell-

based assays for cAMP, intracellular calcium release, and ERK1/2 activation,

we demonstrate the assay-dependence of efficacy and the range of signaling

responses achievable through biased agonism. These results are then dis-

cussed in the context of the known effects of these same ligands in vivo to

illustrate the difficulty and complexity of using in vitro profiles of ligand

activity to predict biological response.

2. DETERMINING THE RELATIVE ACTIVITY OF PTH1RLIGANDS

For any given GPCR ligand, an activity profile can be generated by

determining its EC50 and Emax across a panel of assays measuring different

indices of receptor activation. These results will be specific for each rec-

eptor–ligand combination, but they will also be subject to system factors

influencing coupling efficiency, such as receptor density, that can cause

ligand classification to appear to vary between assays. Such factors cannot

change the relative order of potency or efficacy for a series of ligands, but

can create the appearance of signal bias. For example, a ligand may appear

to be a full agonist in terms of both potency and efficacy if the signal is tightly

coupled, that is, there is significant “receptor reserve,” or highly amplified,

for example, second messenger production, and as a partial agonist when

assaying a response that is weakly coupled, that is, all receptors must be in

the active state to achieve a maximal response, or unamplified, for example,

stoichiometric GPCR–arrestin binding.


When characterizing a series of ligands across multiple assays of receptor

activation, it is useful to determine intrinsic relative activity (RAi). RAi is a

measure of the microscopic affinity constant of an agonist for the active

state of the receptor expressed relative to that of a reference agonist, which

in its simplest form can be estimated from EC50 and Emax values using the

equation (Griffin, Figueroa, Liller, & Ehlert, 2007):

RAi¼Emax�BEC50�A

Emax�AEC50�B

where Emax is the maximum observed system response, EC50 is the ligand

concentration producing a half-maximal response, and A and B are the ref-

erence and test ligands, respectively. This simple calculation of RAi is valid

in cases where the Hill slope of the agonist concentration–response curve is

close to 1.0 or the Emax values of the agonists are equivalent.

When these conditions are not met, cleaner quantification of ligand bias

can be obtained using the Black/Leff operational model (Black & Leff, 1983),

which relates the equilibrium dissociation constant (KA) of the ligand, a direct

measure of receptor occupancy, to a coupling efficiency factor (t), whichencompasses both the intrinsic efficacy of the agonist and system-dependent

factors such as receptor density and coupling efficiency (Kenakin, 2009). Since

the latter factors are constant for any concentration–response curve deter-

mined for any given signaling pathway in the same cell, the ratio of t valuesfor any two agonists in the same system will yield a ratio of intrinsic efficacy

that is independent of receptor number or coupling efficiency. Because allo-

steric effects exerted by other system components can alter ligand affinity as

well as efficacy, it cannot be assumed that the KA value for a given agonist

will be constant under all conditions, so t/KA ratios must be determined for

each assay system. Once determined for each agonist/pathway of interest,

t/KA ratios can be used to quantify bias relative to a reference agonist.

Despite the additional data required, the advantages of the operational model

are its ability to quantify the full range of agonism from submaximal effects to

effects in very sensitive systems with receptor reserve, and that t/KA ratios

determined in one system are applicable to all systems without the need to

individually quantify functional selectivity in all systems. Variation in recep-

tor density and coupling efficiency between systems might change the ability

of all agonists targeting a given receptor to activate a particular pathway, but

it will not change the pathway selective bias of different ligands relative to

one another. While this more generalizable approach has proven useful in

discriminating “weak” ligand bias that might otherwise be obscured by sys-

tem factors (Figueroa et al., 2009; Rajagopal et al., 2011), it remains unclear


whether the added sensitivity of the operational model will prove useful in

industrial applications.

2.1. Defining reference and test ligandsThe PTH1R is a class 2 GPCR highly expressed in kidney and bone that

mediates the effects of PTH, an 84-amino acid peptide hormone that func-

tions as the primary systemic regulator of calcium homeostasis. Most of its

actions are mediated by classic G protein signaling mechanisms, including

Gs-mediated activation of adenylyl cyclase, resulting in cAMP production

and PKA activation, as well as Gq/11- or Gi/o-mediated activation of

phospholipase-Cb, leading to inositol-1,4,5-trisphosphate production, cal-

cium mobilization, and PKC activation (Gesty-Palmer & Luttrell, 2011).

The PTH1R also engages arrestins, which mediate both receptor desensitiza-

tion and arrestin-dependent signaling (Ferrari, Behar, Chorev, Rosenblatt, &

Bisello, 1999; Gesty-Palmer et al., 2006). In transfected HEK293 cells,

PTH1R-mediated ERK1/2 activation results from two temporally distinct

mechanisms: a conventional G protein-dependent pathway that involves

PKAand/orPKCand aGprotein-independent pathwaymediated by arrestins

(Gesty-Palmer et al., 2006).

The PTH1R has long served as a model for the study of functional selec-

tivity inGPCR signaling, as its pleiotropic downstream signaling is sensitive to

changes in ligand structure. Whereas the C-terminal truncated PTH(1–34)

fragment possesses all of the known biochemical and physiologic properties

of the native hormone, acting as a conventional/full agonist with respect to

activation of Gs and Gq/11 signaling and arrestin-dependent receptor desensi-

tization and internalization, other PTH fragments exhibit marked variations in

coupling PTH1R to downstream effectors. For example, shorter N-terminal

fragments of the PTH peptide, for example, PTH(1–31), activate adenylyl

cyclase in ROS 17/2 rat osteosarcoma cell membranes without stimulating

membrane-associated PKC ( Jouishomme et al., 1994), while N-terminal

truncations, for example, PTH(3–34), activate PKC while failing to activate

adenylyl cyclase (Jouishomme et al., 1992). Further N-terminal truncations,

for example, PTH(7–34), which still possess the structural determinants nec-

essary for relatively high affinity binding but lack the N-terminal residues

needed to stimulate guanine nucleotide exchange, antagonize G protein sig-

naling but still stimulate receptor phosphorylation and internalization

(Sneddon et al., 2004). Other signal-selective PTH analogs include Trp1-

PTHrp(1–36), which has been reported to activate ERK1/2 in HEK293 cells


solely through a Gs/PKA-dependent pathway that is unaffected by PKC

inhibition or downregulation of arrestin expression, suggesting that it is

Gs-selective (Gesty-Palmer et al., 2006), and (D-Trp12, Tyr34)-bPTH

(7–34), which acts as an inverse agonist for adenylyl cyclase activation, yet

is capable of activating ERK1/2 via a arrestin-mediated signaling pathway

(Gardella et al., 1996; Gesty-Palmer et al., 2006).

If the goal is to compare the ability of ligands to bias PTH1R signaling,

prior studies are limited by the fact that they were, in most cases, performed

in different cell backgrounds and/or used different assays to measure effector

coupling. To produce directly comparable data, we chose a panel of PTH

analogs based on literature reports (Table 13.1) and compared them in three

assays of PTH1R receptor activation that reflect most of the downstream

Table 13.1 Reported activity profiles of selected PTH1R ligands

Ligand Kd (nM)G-proteincoupling

Arrestincoupling References

PTH(1–34) 2�1 Gs and Gq/11 Arrestin

2/3

Juppner et al. (1991), Abou-Samra

et al. (1992), Bringhurst et al.

(1993), Pines et al. (1994), Takasu,

Guo, and Bringhurst (1999), Gesty-

Palmer et al. (2006)

PTH(1–31) NDa Gs and Gq/11 ND Whitfield and Morley (1995),

Takasu and Bringhurst (1998),

Sneddon et al. (2004)

Trp1PTHrp

(1–36)

ND Gs only Antagonist Gesty-Palmer et al. (2006)

PTH(3–34) 10 Gs and Gq/11

antagonist

ND Segre, Rosenblatt, Reiner,

Mahaffey, and Potts (1979),

Nussbaum, Rosenblatt, and Potts

(1980), Takasu, Guo, et al. (1999)

PTH(7–34) 58�16 Gs and Gq/11

antagonist

ND Hoare and Usdin (2000), Sneddon

et al. (2004)

D-Trp12,

Tyr34-

bPTH(7–

34)

25�2 Gs inverse

agonist

Gq/11

antagonist

Arrestin

2/3

Gardella et al. (1996), Hoare and

Usdin (2000), Gesty-Palmer et al.

(2006), Gesty-Palmer et al. (2009)

ND, not determined.aPTH(1–31) IC50 for PTH(1–34) binding¼78�8 nm (Barbier et al., 2005).


PTH1R actions that are reportedly subject to ligand bias: cAMP production,

intracellular calcium mobilization, and ERK1/2 phosphorylation. All assays

were performed using human PTH1R ectopically expressed in a HEK293

cell background that lacks endogenous PTH1R. For each ligand in each

assay, RAi estimates were generated using human PTH(1–34) as the

reference ligand.

2.2. Assaying hPTH1R-mediated cAMP productionGPCR effects on intracellular cAMP concentration are largely a reflection of

the activation of adenylyl cyclases by Gs and inhibition by Gi/o proteins.

PTH1R effects on cAMP were determined using the Promega GloSensor

cAMP assay, which permits real-time measurement of cAMP concentra-

tion in live cells (Binkowski, Fan, & Wood, 2011). The GloSensor cAMP

reporter is composed of a genetically modified form of Photinus pyralis lucif-

erase fused to a cAMP-binding protein insert. cAMP binding alters the con-

formation of the reporter to increase luciferase activity. To provide adequate

throughput, the GloSensor assay was adapted for use on the FLIPRTETRA

fluorescence imaging plate reader system that enables simultaneous real-time

recording of luciferase activity in 96-well plate format.

2.2.1 Cell culture and transient expression hPTH1R constructsTomaximize assay consistency, cAMPmeasurements were performed using

GloSensor cAMP HEK293 cells supplied by Promega, Inc. that stably

express the reporter. Assays were performed following transient transfection

of hPTH1R expression plasmids. Two hPTH1R constructs were used: wild-

type hPTH1R was used to assess ligand activity in a system with low basal

receptor-catalyzed Gs activity, while the constitutively active H233R mutant

hPTH1R (Gardella et al., 1996) was used to discriminate inverse agonism

from neutral antagonism.

2.2.1.1 Required materialsCell culture and transfection

• Cells: GloSensor cAMP HEK293 cell line (Promega, Inc.)

• Cell growthmedium:Dulbecco’smodifiedEaglesMedium(DMEM) sup-

plemented with 10% fetal bovine serum (FBS), 1% antibiotic–antimycotic

solution, and 50 mg/mL hygromycin B

• cDNA expression plasmids: Wild-type and H223R hPTH1R cDNAs

cloned into the pCMV6-XL6 expression vector (Origene Technologies,

Inc.)


• Transfection reagents and medium: Lipofectamine 2000 Transfection

Reagent (Invitrogen—Life Technologies Corp.); OptiMEM

(Gibco—Life Technologies Corp.); serum-free Minimum Essential

Medium (MEM)

• Low serum medium: MEM supplemented with 1% FBS and 50 mg/mL

hygromycin B

Additional solutions

• Cellgro 0.05% trypsin solution (Mediatech, Inc.)

• Collagen solution: 800 mL of rat tail collagen (Becton, Dickson and

Company) in 49.2 mL of sterile 2% acetic acid in distilled H2O, sterilized

using a 0.2-mm filter sterilization unit

• 1� phosphate-buffered saline (PBS)

Disposables

• 10-cm tissue culture dishes

• 1.5-mL sterile Eppendorf microcentrifuge tubes

• 10-mL sterile culture tubes

• Costar white-walled clear-bottom 96-well plates

2.2.1.2 Culture and transient transfection of GloSensor cAMP HEK293 cellsCell culture. The GloSensor cAMP HEK293 cell line was maintained for up

to 20 passages on 10-cm culture dishes in DMEM growth medium con-

taining 50 mg/mL hygromycin B for selection. Cells were maintained at

37 �C in a 5% CO2 atmosphere and passed by trypsinization every

3–4 days to maintain subconfluence.

Transient transfection of GloSensor cAMPHEK293 cells. On day 1, GloSensor

cAMPHEK293 cellswere split into 10-cm tissue culture dishes at a density suf-

ficient to achieve 50% confluence by day 2. GloSensor cAMP HEK293 cells

were transiently transfected with wild-type or H223R hPTH1R expression

plasmids on day 2. Using sterile Eppendorf tubes, 10 mg of PTH1R or

H223R PTH1R plasmid DNA was added to a final volume of 500 mL of

OptiMEMin tube“A”and2.5 mLof lipofectaminewas added toa final volume

of 500 mL ofOptiMEMin tube “B.”TubesA andBwere vortexed briefly and

left to incubate at room temperature for 5 min. Following incubation, tube B

was added to tube A, inverted twice to mix, and left to incubate at room

temperature for 20 min. The 1 mL mixture of DNA, Lipofectamine, and

OptiMEM was added to a 10-mL conical tube containing 1 mL prewarmed

OptiMEM and 3 mL of serum-free MEM. Following medium aspiration

and 1� PBS wash, the 5 mL mixture was added to each 10-cm plate of

GloSensorcAMPHEK293cells and left to incubate at 37 �Cfor4 h.Following


incubation, the transfection medium was aspirated and replaced with growth

medium, and the plates were returned to the tissue culture incubator.

Preparation of assay plates. Prior to cell plating, sterile Costar white-walled

clear-bottom96-well plateswerecollagencoatedbyadding sufficientcollagen/

acetic acidmixture to cover thebottomof eachwell.Theplateswere incubated

at room temperature for 1 h, after which the collagen solution was aspirated,

and each well was washed twice with 100 mL of 1� PBS. On day 3, the trans-

fected cells in 10-cm plates were trypsinized and seeded into collagen-coated

96-well white-walled clear-bottom plates in growth medium at a density of

5�104 cells/well.On day 4, the growthmediumwas aspirated fromeachwell

and replaced with 1% FBS MEM including 50 mg/mL hygromycin B and

returned to the tissue culture incubator overnight.

2.2.2 GloSensor cAMP assay using the FLIPRTETRA

To measure cAMP in the FLIPRTETRA using GloSensor cAMP HEK293 cells

transfectedwithwild-type orH223Rmutant PTH1R, 96-well plates of trans-

fected cells were pre-equilibrated in GloSensor cAMP reagent. Drug plates

containing serial dilutions of each test ligand at 5� final concentration were

prepared for dispensing by the FLIPRTETRA. Changes in luminescence were

recorded in real-time following the injection of ligand and normalized signal

intensity was used to generate ligand concentration–response relationships.

2.2.2.1 Required materialsInstrumentation

• FLIPRTETRA fluorescence imaging plate reader system (Molecular Dynam-

ics, Inc.). In luminescencemode, excitation andemission filters are disabled

and the emission filter removed to allow direct luminescence detection.

Reagents and materials

• PTH1R test and reference ligands: hPTH(1–34) (Bachem, Inc.); hPTH

(1–31) (Bachem, Inc.); bPTH(3–34) (Bachem, Inc.); hPTH(7–34)

(Bachem, Inc.); Trp1-hPTHrp(1–36) (American Peptide Co.); D-

Trp12, Tyr34-bPTH(7–34) (Bachem, Inc.), dissolved in sterile distilled

H2O at 0.1 mM and 1 mM stock concentrations and aliquoted into single

use volumes to avoid freeze/thaw of the peptides.

• Clear round bottom 96-well plates

• Single and multichannel pipettors


• Promega GloSensor cAMP reagent reconstituted according to the man-

ufacturer’s protocol and stored in 200 ml aliquots at �80 �C until use


• Serum-free MEM supplemented with 10 mM HEPES, pH 7.4

• 10 mM forskolin stock solution in dimethylsulfoxide

• 5 mM 3-isobutyl-1-methylxanthine solution (IBMX) in ethanol

Disposables

• Micropipettor tips and solution dispensing trays

• 10-mL sterile culture tubes

• 1.5-mL Eppendorf microcentrifuge tubes

2.2.2.2 Performing the GloSensor cAMP assayLoading the GloSensor cAMP reagent. cAMP assays were performed on day 5

(72 h after transfection). Fresh cAMP reagent medium was prepared by

adding a freshly thawed aliquot of 200 mL of GloSensor cAMP reagent to

10 mL of serum-free MEM buffered with 10 mM HEPES, pH 7.4. The

growth medium was gently aspirated from assay plates and replaced with

100 mL/well of prewarmed cAMP reagent medium using a multichannel

pipettor. Plates were incubated at 37 �C with 5% CO2 for 1 h and then

removed from the incubator and incubated at room temperature in the dark

for an additional 30 min.

Preparation of the ligand dosing plate. Drug plates for the FLIPRTETRA dis-

pensing system were prepared during the cell preincubation period. Fresh

serial dilutions of each PTH1R ligand at 5� working concentration were

prepared in serum-free MEM buffered with 10 mMHEPES.Working con-

centrations spanned the expected active concentration range of each ligand

(10�11 to 10�5 M depending on the ligand). A round-bottom 96-well drug

plate was designed to contain 40 mL of 5� drug concentration per well.

Each concentration was tested in replicates of three on the plate. 10 mMforskolin, which directly stimulates adenyl cyclase, was included on each

drug plate as a positive control.

GloSensor cAMP assay. Excitation and emission filters were removed

prior to initializing the FLIPRTETRA. All assays were run at room tempera-

ture. The instrument was programmed to dispense a total of 25 mL of vehiclecontrol, ligand, or forskolin from each well of the drug plate into the 100 mLof medium in the corresponding well of the assay plate to reach the working

concentration. For measuring luminescence, detection gain was set to

280,000 with exposure time 0.53 s and the gate open 100%. Assay and ligand

dosing plates were loaded into the instrument and luminescence was

recorded every 1 s for 10 reads to establish baseline luminescence, then every

1 s for 50 reads. Thereafter, luminescence was recorded every 2 s for 600

reads (660 total reads over 21 min). In the FLIPRTETRA, the maximum


change in luminescence was reached approximately 5 min after the addition

of ligand and remained stable for at least 10 min. Raw data representing the

time–luminescence relationship for each well following ligand addition

were exported to Microsoft Excel for background subtraction and analysis.

The relative expression levels of wild-type and H223R mutant PTH1R

were verified to be similar by immunoblotting. The protocols for wild-type

and H223R hPTH1R were identical, except that for H223R hPTH1R

experiments, IBMX, a nonselective phoshosdiesterase inhibitor, was used

to improve assay sensitivity for detecting inverse agonism. 1 mL of 5 mM

IBMXwas added to each well for the final 10 min of room temperature pre-

incubation in cAMP reagent medium prior to loading the assay plates into

the instrument.

2.2.3 Estimating RAi for PTH1R-mediated cAMP productionFor each ligand, normalized concentration–response curves were generated

from the raw cAMP luminescence data. GraphPad Prism software was used

to calculate Emax, EC50, and Hill slope. RAi was estimated from measured

Emax and EC50 values using PTH(1–34) as the reference ligand.

2.2.3.1 Required materialsComputer software

• Microsoft Excel

• GraphPad Prism

2.2.3.2 Data processing and resultsGenerating ligand concentration–response curves. Measurements were taken from

each well after stable maximum luminescence was attained. UsingMicrosoft

Excel, background luminescence measured in vehicle-treated wells was sub-

tracted from maximal luminescence in ligand-treated wells to yield the net

change in luminescence. The mean net change in luminescence from trip-

licate wells at each ligand concentration was determined, and all values were

normalized to the peak luminescence observed with PTH(1–34). Using

GraphPad Prism, each normalized concentration–response dataset was fit

to a sigmoidal dose–response curve using a variable Hill slope. Emax and

EC50 were determined from these curves. A minimum of three separate

experimental replicates were performed using each ligand.

As shown in Fig. 13.1, experiments performed using wild-type hPTH1R

clearly separated ligands into two groups. hPTH(1–34), hPTH(1–31), and

Trp1-hPTHrp(1–36), all appeared as full agonists, producing a similar

Ligand EC50 Emax RAiPTH (1–34) 1.85 � 10-8

2.79 � 10-8

2.92 � 10-8

1.0

PTH (1–34)

PTH (1–31)

(D-Trp12, Tyr34

)-PTH (7–34)

log, [Ligand] M

-100.0

0.2

0.4

0.6

0.8

1.0

1.2

-8 -6 -4

% M

ax P

TH

(1–

34)

resp

onse

PTH (3–34)

PTH (7–34)

(Trp1)-PTHrp (1–36)

1.0

PTH (1–31) 1.074 0.71

(Trp1)-PTHrp (1–36) 0.94 0.64

PTH (3–34) N.D. N.S. 0

PTH (7–34) N.D. N.S. 0

(D-Trp12,Tyr34)-PTH (7–34) N.D. N.S. 0

A

B

Figure 13.1 Ligand-mediated cAMP production by wild-type PTH1R. (A) Ligandconcentration–response relationships were determined for a panel of six PTH1R peptideligands using GloSensor HEK293 cells transiently overexpressing hPTH1R as described inthe text. The mean net change in cAMP reporter luminescence was normalized to themaximal net response elicited by the reference ligand, PTH(1–34). Sigmoidal dose–response curves and SEM across at least three independent experiments were gener-ated using GraphPad Prism. (B) For each ligand, RAi was estimated from the observedEmax and EC50 values. Ligands that failed to generate a statistically significant change incAMP luminescence were assigned a RAi of zero. NS, no significant response; ND, notdetermined.


maximal response. In contrast, bPTH(3–34), hPTH(7–34), and D-Trp12,

Tyr34-bPTH(7–34) produced no significant change from basal. Figure 13.2

depicts results obtained using the H223R hPTH1R. Normalized basal

cAMP luminescence in cells expressing the H223R hPTH1R was about

0.28 of the maximal PTH(1–34)-induced luminescence observed with

the wild-type hPTH1R, reflecting the constitutive activity of the mutant.

When ligand-induced changes in cAMP luminescence obtained in cells

expressing H223R hPTH1R are normalized to this higher basal, the

concentration–response curves clearly distinguish D-Trp12, Tyr34-bPTH

(7–34) as an inverse agonist for cAMP production, while bPTH(3–34)

Ligand EC50 Emax RAiPTH (1–34) 8.40 � 10–9

1.26 � 10-8

1.79 � 10-8

3.36 ´ 10-9

1.0

PTH (1–34)

PTH (1–31)

PTH (3–34)

PTH (7–34)

(D-Trp12,Tyr34)-PTH (7–34)

log, [Ligand] M

% N

orm

aliz

ed m

ax P

TH

(1–

34)

resp

onse

-10-0.6

-0.4

-0.2

0.0

0.2

0.4

0.6

0.8

1.0

1.2

-8 -6 -4


1.0

PTH (1–31) 1.07 0.69

(Trp1)-PTHrp (1–36) 0.85 0.57

PTH (3–34) N.D. N.S. 0

PTH (7–34) N.D. N.S. 0

(D-Trp12,Tyr34)-PTH (7–34) -0.44 -1.12

A

B

Figure 13.2 Ligand-mediated cAMP production by the constitutively active H223Rmutant PTH1R. (A) Ligand concentration–response relationships were determined forthe PTH1R ligand panel using the same protocol described in Fig. 13.1, except thatGloSensor HEK293 cells were transfected with the H223R-mutant PTH1R, which gener-ates constitutive receptor-mediated cAMP production, permitting detection of inverseagonism. Ligand effects are depicted as positive or negative change from basal cAMPluminescence in the constitutively active system. (B) RAi estimates reflect the directionof change in cAMP luminescence relative to the reference agonist. The negative RAi cal-culated for (D-Trp12, Tyr34)-bPTH (7–34) reflects its inverse agonism of receptor–Gs cou-pling. NS, no significant response; ND, not determined.


and hPTH(7–34) remain neutral. These results are consistent with literature

reports of the effects of each ligand on PTH1R-Gs coupling (Gardella et al.,

1996; Gesty-Palmer et al., 2006; Jouishomme et al., 1992, 1994; Sneddon

et al., 2004).

Estimating RAi for cAMP production. Because the empirically determined

Hill slopes for PTH1R ligand effects on cAMP production were close to 1.0,

RAi values were estimated using the simplified formula given in Section 2.

Emax/EC50 ratios for each ligand were normalized to that of the reference

ligand, PTH(1–34) (Figs. 13.1 and 13.2). For cAMP production determined

using the wild-type PTH1R, the activity rank order was hPTH(1–34)


¼hPTH(1–31)¼Trp1-hPTHrp(1–36)>>>bPTH(3–34)¼hPTH(7–34)

¼D-Trp12, Tyr34-bPTH(7–34). Using the H223R PTH1R, the observed

activity rank order was hPTH(1–34)>hPTH(1–31)¼Trp1-PTHrp

(1–36)�bPTH(3–34)¼hPTH(7–34)�D-Trp12, Tyr34-bPTH(7–34).

2.3. Assaying PTH1R-mediated intracellular calcium influxGPCR effects on intracellular calcium arise largely from activation of

phospholipase-Cb isoforms by Gq/11 and Gbg subunits primarily derived

from Gi/o proteins. The PTH1R regulates phospholipase-Cb through cou-

pling to both Gq/11 and Gi/o in varying proportions depending on cell back-

ground. In osteoblastic cells, PTH-stimulated phospholipase-Cb activation is

primarily Gq/11-dependent, while renal tubular cells exhibit Gi/o-dependent

phospholipase-Cb activation due to expression of the Naþ/Hþ exchanger

regulatory factors 1 and 2, which enhance PTH1R-Gi/o coupling (Mahon,

Donowitz, Yun, & Segre, 2002). Because measurement of intracellular

calcium using calcium-sensitive fluorescent dyes is readily amenable to

high-throughput screening, we chose to assay ligand-induced calcium flux

as a marker for PTH1R regulation of the phospholipase-Cb–inositol-1,4,5-trisphosphate–intracellular calcium–PKC pathway. While this assay predom-

inantly reflects phospholipase-Cb activation in short-term stimulations, it does

not discriminate between the contributions of Gq/11 and Gi/o proteins and

thus is unable to distinguish possible mechanistic ligand bias between these

two effectors.

2.3.1 Cell culture and inducible expression of hPTH1RTo maximize consistency between experiments, calcium assays were per-

formed using a stable HEK293-FlpIn TRex cell line engineered for

tetracycline-inducible expression of wild-type hPTH1R. Assays were

performed following the induction of hPTH1R expression.

2.3.1.1 Required materials• Cells: Tetracycline-inducible hPTH1RHEK293-FlpIn TRex cells were

established by cloning cDNA encoding the hPTH1R into the HEK293-

FlpIn TRex cell line using previously described methods (Zimmerman,

Simaan, Lee, Luttrell, & Laporte, 2009)

• Cell growth medium: Phenol red-free DMEM supplemented with 10%

FBS, 1% antibiotic–antimycotic solution, and 50 mg/mL hygromycin B

plus 5 mg/mL blasticidin for selection


• Serum-free medium: Phenol red-free DMEM supplemented with 0.05%

bovine serum albumin



• 1 mg/mL tetracycline stock solution in sterile distilled H2O

Disposables


• Costar 96-well black-wall clear-bottom plates

2.3.1.2 Cell culture and induction of PTH1R expressionCell culture. hPTH1R HEK293-FlpIn TRex cells were maintained for up to

20 passages on 10-cm culture dishes in phenol red-free DMEM growth

medium containing 50 mg/mL hygromycin B plus 5 mg/mL blasticidin

for selection. The cells were maintained at 37 �C in a 5% CO2 atmosphere

and passed by trypsinization every 3–4 days to maintain subconfluence.

Tetracycline induction of hPTH1R expression. On day 1, hPTH1R HEK293-

FlpIn TRex cells were seeded into dishes at a density sufficient to attain 50%

confluence by day 2. At the time of passage, 1 mg/mL tetracycline was added

to the growth medium to induce PTH1R expression. The cells remained in

tetracycline-supplemented medium until assay 72 h later.

Preparation of assay plates. On day 2, the cells were seeded at a density of

5�104 cells/well into black-wall clear-bottom 96-well plates that were pre-

coated with collagen as described in Section 2.2.1.2 and cultured in growth

medium containing tetracycline, hygromycin B, and blasticidin. On day 4,

the growth medium was aspirated and the cells were serum-starved for 3 h

prior to assay in 100 mL of phenol red-free DMEM supplemented with

0.05% BSA and tetracycline.

2.3.2 FLIPRTETRA calcium assayMeasurements of ligand-induced calcium flux were performed on the

FLIPRTETRA fluorescence imaging plate reader system using the FLIPR

Calcium Assay Kit 5 from Molecular Devices, Inc. Tetracycline-induced

hPTH1R HEK293-FlpIn TRex cells in 96-well plates were preincubated

with a proprietary calcium-sensitive fluorescent dye while drug plates

containing serial dilutions of the test ligands were prepared. Changes in

dye fluorescence were recorded in real-time following the injection

of ligand and normalized signal intensity was used to generate agonist

concentration–response relationships.


2.3.2.1 Required materialsInstrumentation

• FLIPRTETRA fluorescence imaging plate reader system (Molecular

Dynamics, Inc.) with 470–495 nm excitation light-emitting diodes

and 515–575 nm emission filter


• PTH1R test and reference ligands

• FLIPR Calcium 5 Assay Kit (Molecular Devices, Inc.)

• Clear round-bottom 96-well plates

• Single and multichannel pipettors


• 10 mM A23187 stock dissolved in dimethylsulfoxide

• 1� PBS

Disposables

• Micropipettor tips and solution dispensing trays


2.3.2.2 Performing the fluorescence-based calcium assay using the FLIPRTETRA

Loading the calcium indicator dye. The calcium indicator dye was prepared

according to manufacturer’s protocol and 100 mL was added to the 100 mLof starvation medium in each well for a final volume of 200 mL/well. Cellswere then incubated for 1 h at 37 �C 5% CO2.

Preparation of the ligand dosing plate. During the preincubation, 96-well

clear round-bottom drug plates for the calcium assay were prepared as

described in Section 2.2.2.2 with the exceptions that ligand dilutions were

performed in 1� PBS and a total of 80 mL of 5� drug concentration was

placed in each well. 10 mM of the calcium ionophore A23187 was used as

the positive control on each drug plate.

Performing the calcium assay. 470–495 nm excitation and 515–575 nm

emission filters were installed prior to initializing the FLIPRTETRA. All assays

were run at room temperature. The instrument was programmed to simul-

taneously dispense 50 mL of vehicle control, 5� ligand, or A23187 from the

drug plate into the 200 mL of medium in the corresponding wells of the assay

plate to achieve the final ligand concentration. For measuring calcium fluo-

rescence, excitation intensitywas set to 50%, and detection gainwas set to 2000

with exposure time 0.53 s and the gate open 10.08%. Assay and ligand dosing

plates were loaded into the instrument and fluorescencewas recorded every 1 s

for 10 reads to establish baseline fluorescence, then every 1 s for 300 reads

(310 total reads over 5.17 min). In the FLIPRTETRA, the maximum change in


fluorescence was attained approximately 50 s after the addition of ligand

and returned to baseline within approximately 150 s. Raw data representing

the time–fluorescence relationship for each well following ligand addition

were exported to Microsoft Excel for background subtraction and analysis.

2.3.3 Estimating RAi for PTH1R-mediated calcium signalingSimilar to theprocess described inSection2.2.3, background fluorescencemea-

sured in vehicle-treated wells was subtracted from peak fluorescence in each

ligand-treated well to yield the net change in fluorescence. The mean net

change in fluorescence from triplicate wells at each ligand concentration

was determined, and all values were normalized to the peak fluorescence

observed with PTH(1–34). Using GraphPad Prism, each normalized

concentration–responsedatasetwas fit to a sigmoidal dose–responsecurveusing

variableHill slope.Emax andEC50valuesweredetermined fromthesecurves.At

least three separate experimental replicates were performed using each ligand.

As shown in Fig. 13.3, only two of the test ligands generated a significant

increase in intracellular calcium in our assay; hPTH(1–34) and hPTH(1–31).

Both ligands demonstrated equivalent efficacy, and although each was

approximately 20-fold less potent in the calcium assay than the cAMP assay,

hPTH(1–34) remained 1.5� more potent than hPTH(1–31). The finding

that hPTH(1–31) stimulates calcium flux via hPTH1R in our HEK293 cell

background probably reflects a tissue difference in hPTH1R signaling.

Although PTH(1–31) is incapable of activating membrane-associated

PKC in ROS 17/2 rat osteosarcoma cells ( Jouishomme et al., 1994), it

robustly elevates intracellular inositol phosphate levels in murine distal con-

voluted tubule cells (Sneddon et al., 2004). Trp1-hPTHrp(1–36) demon-

strated strong bias toward Gs coupling, as it had no significant effect on

intracellular calcium entry despite its ability to stimulate cAMP production

with potency and efficacy equivalent to PTH(1–31). This is consistent with

literature reports of the coupling selectivity of both Trp1-hPTHrp(1–36)

and a related Bpa1-PTHrp(1–36) peptide (Bisello, Horwitz, & Stewart,

2004; Gesty-Palmer et al., 2006). (D-Trp12,Tyr34)-PTH(7–34) also demon-

strated apparent bias in that its inverse agonism of hPTH1R-Gs coupling was

not evident toward hPTH1R-calcium signaling. This too is consistent with

literature reports (Gardella et al., 1996; Gesty-Palmer et al., 2006). As in the

cAMP assay, both hPTH(7–34) and bPTH(3–34) failed to stimulate calcium

flux. The failure of bPTH(3–34) to signal may reflect a species or tissue dif-

ference in PTH1R signaling. Although PTH(3–34) was reported to selec-

tively activate PKC in murine osteosarcoma cells ( Jouishomme et al.,

1992), amino terminal truncation of hPTH was found to completely


6.37 � 10-7

1.0

PTH (1–34)

PTH (1–31)

PTH (3–34)

PTH (7–34)

(D-Trp12,Tyr34)-PTH (7–34)

log, [Ligand] M

% M

ax P

TH

(1–

34)

resp

onse

-70.0

0.2

0.4

0.6

0.8

1.0

1.2

-6 -5


1.0

PTH (1–31) 0.92 0.67

(Trp1)-PTHrp (1–36) N.D. N.S. 0

PTH (3–34) N.D. N.S. 0

PTH (7–34) N.D. N.S. 0

0(D-Trp12,Tyr34)-PTH (7–34) N.D. N.S.

A

B

Figure 13.3 Ligand-mediated stimulation of intracellular calcium flux by the hPTH1R.(A) Ligand concentration–response relationships were determined for the hPTH1R ligandpanel using tetracycline-inducible hPTH1R HEK293-FlpIn TRex cells as described in thetext. The mean net change in calcium fluorescence was normalized to the maximalnet response elicited by the reference ligand, PTH(1–34). Sigmoidal dose–response curvesand SEM across at least three independent experiments were generated using GraphPadPrism. (B) For each ligand, RAi was estimated from the observed Emax and EC50 values.Ligands that failed to generate a statistically significant change in calcium fluorescencewere assigned a RAi of zero. NS, no significant response; ND, not determined.


eliminate hPTH1R stimulation of inositol phosphate hydrolysis in renal

epithelial cells (Takasu, Gardella, Luck, Potts, & Bringhurst, 1999).

As with the cAMP assay, RAi values were estimated using the simplified

formula in Section 2 (Fig. 13.3). For intracellular calcium flux, the activity

rank order was hPTH(1–34)>hPTH(1–31)>>>Trp1-PTHrp(1–36)¼bPTH(3–34)¼hPTH(7–34)¼D-Trp12, Tyr34-bPTH(7–34).

2.4. Assaying PTH1R-mediated ERK1/2 activationRegulation of the ERK1/2 cascade is thought to contribute to the biological

actions of PTH1R, affecting diverse processes such as cellular mitogenesis,

embryologic development, and renal tubular phosphate transport

(Lederer, Sohi, & McLeish, 2000; Verheijen et al., 1999). PTH has been


reported both to activate ERK1/2 in opossum kidney cells (Lederer et al.,

2000) and to inhibit it in rat osteosarcoma cells (Verheijen & Defize, 1995).

Much of this complexity arises from the fact that ERK1/2 represents a

convergence point where multiple upstream signals are integrated. Recent

efforts to deconvolute the hPTH1R-ERK regulatory network in HEK293

cells and murine calvarial osteoblasts suggest that the pathway is indepen-

dently regulated by conventional G protein-dependent pathways involving

Gs-PKA and/or Gq/11-PKC and a G protein-independent pathway medi-

ated by arrestins (Gesty-Palmer et al., 2006, 2009). Evidence suggests that

these different mechanisms control the formation of spatially, temporally,

and functionally discrete pools of ERK1/2 that determine its ultimate bio-

logical function (Luttrell & Gesty-Palmer, 2010).

Largely because ERK1/2 is activated by a multitude of upstream signals,

it is often used for ligand screening.While it offers a single integrated readout

of receptor activation, it is often necessary to perform additional experiments

using pharmacologic inhibitors or RNA interference to determine the con-

tribution of different effectors, for example, arrestins, to the observed signal

(Gesty-Palmer et al., 2006). In addition, G protein-dependent ERK1/2 acti-

vation signals are often highly amplified, while arrestin-dependent ERK1/2

activation, which relies on the assembly of stoichiometric GPCR–arrestin

“signalsomes,” is unamplified, causing arrestin-dependent agonists to have a

reduced apparent efficacy.

2.4.1 Cell culture and inducible expression of hPTH1RERK1/2 assays were performed using the stable hPTH1R HEK293-FlpIn

TRex cell line. In some assays, the arrestin dependence of hPTH1R-mediated

ERK1/2 activation was assessed by determining the loss of signal resulting

from downregulation of arrestin2/3 expression by RNA interference.

2.4.1.1 Required materials• Cells: Tetracycline-inducible hPTH1R HEK293-FlpIn TRex cells

• Cell growth medium: DMEM supplemented with 10% FBS, 1%

antibiotic–antimycotic solution, and 50 mg/mL hygromycin B plus

5 mg/mL blasticidin for selection

• Serum-free medium: DMEM supplemented with 0.05% bovine serum

albumin

• Double-stranded siRNA oligomers (Qiagen, Inc.): Arrestin2/3: 50-AAACCTGCGCCTTCCGC TATG-30; Scrambled control: 50-AAUUCUCCGAACGUGUCACGU-30


• GeneSilencer™ transfection reagent (Gene Therapy Systems)



• 1 mg/mL tetracycline stock solution in sterile distilled H2O

Disposables


• 12-well tissue culture plates

2.4.1.2 Cell culture and induction of PTH1R expressionCell culture. hPTH1R HEK293-FlpIn TRex cells were maintained in growth

medium without tetracycline as described in Section 2.3.1.2 until they were

ready for assay plating.

Induction of hPTH1R expression and arrestin expression silencing. On day 1,

hPTH1R HEK293-FlpIn TRex cells were plated on 10-cm dishes in

growth medium containing 0.1 mg/mL tetracycline to induce PTH1R

expression. On day 3, 6�105 cells/well were seeded into 12-well plates

to establish a confluent monolayer for assay on day 4. On day 4, the growth

medium was aspirated and the cells were serum-starved for 3 h prior to assay

in 1 mL of DMEM supplemented with 0.05% BSA and tetracycline.

For experiments involving downregulation of arrestin2/3 expression by

RNA interference, 50% confluent hPTH1R HEK293-FlpIn TRex cells in

10-cm dishes were transfected on day 2 using 20 mg of scrambled control or

arrestin2/3-targeted siRNA oligomers using 50 ml of GeneSilencer transfec-tion reagent according to the manufacturer’s instructions (Lee et al., 2008).

The cells were seeded in 12-well tissue culture plates on day 3 and serum-

starved for assay on day 4 as described above. Silencing of arrestin expression

was confirmed by immunoblotting whole-cell lysates using rabbit polyclonal

anti-arrestin2/3 IgG with horseradish peroxidase-conjugated donkey anti-

rabbit IgG as the secondary antibody.

2.4.2 Assaying ERK1/2 phosphorylationLigand concentration–response curves were generated by immunoblotting

phospho-ERK1/2 in whole-cell lysates of hPTH1R HEK293-FlpIn TRex

cells following treatment with vehicle or PTH1R ligands.

2.4.2.1 Required materialsEquipment

• Invitrogen XCell SureLock Mini-Cell Electrophoresis System

• Invitrogen iBlot and iBlot nitrocellulose gel transfer stacks

• X-ray film developer

• Kodak Image Station for densitometric image analysis



• PTH1R test and reference ligands

• BCA protein assay kit (Promega)

• Invitrogen NuPAGE Novex precast SDS-PAGE 15-well 4–12% gradi-

ent gels

• Nitrocellulose membranes

• Rabbit polyclonal anti-phosphoERK1/2 IgG (Cell Signaling

Technology)

• Horseradish peroxidase-conjugated goat anti-rabbit IgG (Cell Signaling

Technology)

• SuperSignal West Pico ECL (Promega)

• X-ray film (Kodak)

• X-ray film cassettes


• 200 mM phorbol myristate acetate (PMA) in dimethylsulfoxide

• 1� SDS lysis buffer: 62.5 mM Tris–HCl (pH 6.8), 2% (w/v) SDS, 10%

glycerol, 50 mM DTT, 0.01% (w/v) bromophenol blue

• Tris-buffered saline with Tween 20 (TBST): 20 mM Tris–HCl, pH 8;

150 mM NaCl; 0.001% Tween 20 in distilled H2O.

• Blocking buffer: 5% nonfat dry milk in TBST

• Primary antibody buffer: 5% BSA in TBST

Disposables


• Cell scrapers

• Micropipettor tips

2.4.2.2 Measuring PTH1R-regulated ERK1/2 phosphorylationCell stimulation. Serial dilutions of PTH1R ligands were prepared at 1000�desired final concentration. At time 0, 1 mL of vehicle or ligand concentrate

was added to the 1 mL of serum-free media in each well and the plates were

incubated for 5 min at 37 �C. PMA at a final concentration of 200 nM was

added to one well of each plate as a positive control.

Phospho-ERK1/2 Immunoblotting. Five minutes after ligand application,

the medium was aspirated and monolayers were lysed in 100 mL of 1�SDS lysis buffer, harvested in 1.5-mL Eppendorf tubes by scraping, briefly

sonicated, and clarified by microcentrifugation. The protein content of each

sample was determined using a BCA protein assay kit according to manu-

facturer’s protocol. Equal amounts of protein sample (10 mg) were loaded


onto 4–12% SDS-PAGE gradient gels and proteins resolved by electropho-

resis at 120 V constant voltage for 2 h. The proteins were transferred to

nitrocellulose membranes using the iBlot apparatus according to manufac-

turer’s protocol. The membranes were blocked in 5% dry milk in TBST for

1 h at room temperature with gentle shaking and then incubated overnight

at 4 �C with rabbit polyclonal anti-phosphoERK1/2 antibody at 1:1000

dilution in 5% BSA in TBST with gentle shaking. Following 3�5 min

TBST washes, the membranes were incubated for 1 h at room temperature

with HRP-conjugated goat anti-rabbit IgG at 1:10,000 dilution in 5% dry

milk in TBST with gentle shaking. The membranes were again washed

3�10 min in TBST, developed using SuperSignal West Pico ECL reagent

according to manufacturer’s directions, and exposed to Kodak X-ray film.

Exposure times were varied to capture the widest possible range of band

intensities. Quantification of phosphoERK1/2 pixel intensity was per-

formed using a Kodak Image Station.

2.4.3 Estimating RAi for PTH1R-mediated ERK1/2 activationUsing Microsoft Excel, the net change in phospho-ERK1/2 band intensity

from basal was determined at each ligand concentration, and all values were

normalized to the peak band intensity observed with PMA.Using GraphPad

Prism, each normalized concentration–response dataset was fit to a sigmoidal

dose–response curve using variable Hill slope. Emax and EC50 values were

determined from these curves. At least three separate experimental replicates

were performed using each ligand.

As shown in Fig. 13.4, all the PTH1R ligands except hPTH(7–34) gen-

erated a statistically significant increase in ERK1/2 phosphorylation. All

three ligands that were active in the cAMP assay, hPTH(1–34), hPTH

(1–31), and Trp1-hPTHrp(1–36), produced robust ERK1/2 activation with

nanomolar EC50 values. bPTH(3–34), which was without detectable activ-

ity in either the cAMP or calcium assays, also robustly activated ERK1/2.

The Emax of (D-Trp12,Tyr34)-bPTH(7–34), which demonstrated inverse

agonism for cAMP production and was inactive in the calcium assay, was

less than 10% that of PTH(1–34). The ligands that activated ERK1/2 with-

out producing detectable G protein signals were 3- to 12-fold less potent

than PTH(1–34). hPTH(7–34) appeared as a neutral antagonist in all assays,

failing to elicit measureable changes in cAMP production, intracellular

calcium flux, or ERK1/2 activation.

To estimate the contribution of arrestin-dependent signaling to the

ERK1/2 response, single concentration assays (1 mM) were performed in


PTH (1–34)

PTH (1–31)

PTH (3–34)


(D-Trp12,Tyr34)-PTH (7–34)

log, [Ligand] M

-120.0

0.2

0.4

0.6

0.8

1.0

0.00-10 -8 -6

0.02

0.04

0.06

0.08

0.10

-10 -8 -6 -4

% P

MA

max

res

pons

e

% P

MA

max

res

pons

e

log, [Ligand] M

PTH (7–34)

1.08 � 10-8

4.36 � 10-8

3.42 � 10-9

8.12 � 10-9

0.91 1.0

PTH (1–31) 0.81 0.41

(Trp1)-PTHrp (1–36) 0.85 1.02

PTH (3–34) 0.68 0.26

PTH (7–34) N.D. N.S. 0

(D-Trp12,Tyr34)-PTH (7–34) 0.07 0.007

A

B

Figure 13.4 Ligand-mediated ERK1/2 activation by the hPTH1R. (A) Ligand concentra-tion–response relationships were determined for the hPTH1R ligand panel usingtetracycline-inducible hPTH1R HEK293-FlpIn TRex cells as described in the text. Thenet change in ERK1/2 was normalized to the net response elicited by a maximally effi-cacious dose of PMA. Sigmoidal dose–response curves and SEM across at least threeindependent experiments were generated using GraphPad Prism. (B) For each ligand,RAi was estimated from the observed Emax and EC50 values. Ligands that failed to gen-erate a statistically significant change in ERK1/2 phosphorylation were assigned a RAi ofzero. NS, no significant response; ND, not determined.


hPTH1R HEK293-FlpIn TRex cells after arrestin2/3 expression was

downregulated >80% by RNA interference. Figure 13.5 depicts the sensi-

tivity of basal- and ligand-stimulated ERK1/2 activation to arrestin silenc-

ing. Consistent with the literature reports suggesting that arrestin signaling

contributes to ERK1/2 activation by conventional ligands (Ahn, Shenoy,

Wei, & Lefkowitz, 2004; Gesty-Palmer et al., 2006), hPTH(1–34) and

hPTH(1–31) responses were modestly attenuated, demonstrating approxi-

mately 20% reduction of the peak ERK1/2 activation observed at 5 min

stimulation. In contrast, ERK1/2 activation by bPTH(3–34) and

(D-Trp12,Tyr34)-bPTH(7–34), which showed antagonist or inverse agonist

activity in assays of G-protein signaling, were 70–80% sensitive to arrestin

siRNA Arrestin 2/3

pERK IB

Scrambled

NS

% I

nhib

itio

n of

ER

Kph

osph

oryl

atio

n

PTH (1

–34)

0

20

40

60

80

100+ -- +

+ -- +

+ -- +

+ -- +

+ -- +

+ -- +

PTH (1

–31)

PTH (3

–34)

(D-T

rp12 ,T

yr34 )-P

TH (7

–34)

(Trp1 )-P

THrp (1

–36)

Figure 13.5 Sensitivity of ligand-mediated ERK1/2 activation to downregulation ofarrestin2/3 expression. To elucidate the arrestin dependence of ERK1/2 activation bydifferent hPTH1R ligands, hPTH1R HEK293-FlpIn TRex cells were transfected with scram-bled control or arrestin2/3-targeted siRNA prior to determining the ERK1/2 response to amaximally effective concentration (1 mM) of each ligand that produced significant ERK1/2activation. The upper panel shows a representative immunoblot taken from one of threeindependent experiments. The bar graph presents the percentage inhibition of thephosphoERK1/2 signal in arrestin2/3 siRNA-treated cells normalized to the responsein control siRNA-treated cells. As shown, ligands that stimulated ERK1/2 withoutdetectably stimulating cAMP production were more sensitive to arrestin silencing thanthose producing a robust G protein-mediated response.


downregulation. Interestingly, Trp1-PTHrp(1–36), which has been described

as a “nondesensitizing” Gs-selective hPTH1R ligand, was intermediate in its

sensitivity to arrestin knockdown. Overall, these data are consistent with

reports that arrestin-dependent signaling accounts for the majority of G

protein-independent ERK1/2 activation by GPCR ligands (Luttrell &

Gesty-Palmer, 2010).

As with the preceding assays, RAi values were estimated using the sim-

plified formula in Section 2. For ERK1/2 activation, the activity rank order

was hPTH(1–34)>Trp1-PTHrp(1–36)>hPTH(1–31)>bPTH(3–34)>D-Trp12, Tyr34-bPTH(7–34)>hPTH(7–34). Because (D-Trp12,Tyr34)-

bPTH(7–34) exhibited less than 1/10th the apparent efficacy and a 12-fold

lower potency than PTH(1–34), the estimated RAi for ERK1/2 activation

is markedly lower than that for the other active ligands.


2.5. Estimating PTH1R ligand biasFigure 13.6A presents a simple graphic representation of the activity profiles of

our ligand panel based onRAi estimates. As shown, hPTH(1–34) behaves as a

full agonist in all three assays, while hPTH(1–31) appears as a relatively

A BpERK1/2

pERK1/2

Ca2+i–1.0

–0.8

–0.6

–0.4

–0.2

–0.2 0.2 0.4 0.6 0.8 1.0–0.4–0.6–0.8–1.0 0

–0.2

0

0.2

0.4

0.6

0.8

1.0

–0.4

–0.6

–0.8

–1.0

Ca2+i

cAMP cAMP

0.2

1.0

0.8

0.6

0.4pER

K

0.2

1.0

D C

0.8PTH(1–34)PTH(1–31)(Trp1)-PTHrp (1–36)PTH (3–34)PTH (7–34)(D-Trp12,Tyr34)-PTH (7–34)

0.6

0.4pER

K

0.2

0.00.0 0.2 0.4

Ca2+0.6 0.8 1.0 –1.0 –0.5 0.0

cAMP

0.5 1.0

–1.0 –0.5 0.0

cAMP

0.5 1.0

0.4

Ca2

+

0.6

0.8

1.0

0.2

0.4

0.6

0.8

Figure 13.6 Graphic analysis of PTH1R ligand bias. (A) Multiaxial representation of PTH1Rligand activity in the cAMP, calcium, and ERK1/2 assays. Estimated RAi values for eachligand are plotted on each axis to represent themagnitude and direction of effect in eachsignaling response. Ligands with “balanced” efficacy, such as hPTH1-34 and hPTH(1–31),show effects of similar amplitude and direction on all three axes, while those demonstrat-ing “bias” show disproportionate activity in one or more pathways or reversal of efficacy.(B–D) Pairwise comparison of ligand activity at equimolar concentration in cAMP versuscalcium signaling (B), cAMP versus ERK1/2 phosphorylation (C), and calcium signaling ver-sus ERK1/2 phosphorylation (D). In each plot, deviation from the line of unity reflects dif-ferences in coupling efficiency between the receptor and its downstream effectors, forexample, cAMP and calcium signaling. Ligands whose coupling efficiency between effec-tor pathwaysmatches that of the reference ligand, for example, PTH(1–34) and PTH(1–31),are unbiased, while those that deviate significantly exhibit functional selectivity. (D-Trp12,Tyr34)-bPTH(7–34), which acts as an inverse agonist for cAMP production, is neutral forcalcium signaling and is a partial agonist for ERK1/2 activation, demonstrates assay-dependent reversal of efficacy, a hallmark of biased GPCR agonism.


“balanced” partial agonist. hPTH(7–34), which was inactive in all assays,

appears as a “neutral” antagonist. The other three ligands exhibit apparent sig-

nal bias. Trp1-PTHrp(1–36) appears to be selective for Gs-cAMP signaling,

since it fails to elicit a threshold calcium signal at concentrations that robustly

activate cAMP production and ERK1/2, whereas hPTH(1–34) and hPTH

(1–31) exhibit proportional RAi in all three assays. bPTH(3–34) exhibitsmore

drastic bias, retaining substantial activity in the ERK1/2 assay in the absence of

a detectable cAMP or calcium signal. (D-Trp12,Tyr34)-bPTH(7–34) exhibits

frank reversal of efficacy, one of the hallmarks of biased agonism, appearing as

an inverse agonist for cAMP production, neutral for calcium signaling, and a

partial agonist for ERK1/2 activation.

A useful way of visualizing ligand bias is to plot the response observed in

two different assays at equal ligand concentrations against one another

(Gregory, Hall, Tobin, Sexton, & Christopoulos, 2010). The resultant plot

is a direct comparison of the efficiency of signaling through the two path-

ways. Figure 13.6B shows the relationship between cAMP and calcium sig-

naling for our panel of six PTH1R ligands. In our HEK293 cell background,

the reference ligand hPTH(1–34) is more efficiently coupled to cAMP sig-

naling, in that a significant increase in cAMP production occurs before the

rise in the calcium response, that is, the calcium concentration–response

curve is right-shifted relative to the cAMP curve. hPTH(1–31) appears as

a balanced agonist, in that its cAMP-calcium signal coupling overlaps that

of the reference ligand. bPTH(3–34) and hPTH(7–34) appear as balanced

antagonists, producing no significant change in either assay. Both Trp1-

hPTHrp(1–36) and (D-Trp12,Tyr34)-bPTH(7–34) show apparent signal bias.

Trp1-hPTHrp(1–36) exhibits Gs-selective agonist bias, in that it achieves

cAMP production activity equivalent to the reference agonist while failing

to significantly increase calcium flux. Similarly, (D-Trp12,Tyr34)-bPTH

(7–34) exhibits Gs-selective inverse agonist bias contrasting with neutral

antagonism of calcium signaling.

Figure 13.6C plots the relationship between cAMP and ERK1/2 activa-

tion. Here, the strongly Gs-coupled ligands, hPTH(1–34), hPTH(1–31), and

Trp1-hPTHrp(1–36) appear unbiased, and hPTH(7–34) remains a neutral

antagonist. Both bPTH(3–34) and (D-Trp12,Tyr34)-bPTH(7–34) show signal

bias. bPTH(3–34) robustly activates ERK1/2 without producing a significant

cAMP signal, while (D-Trp12,Tyr34)-bPTH(7–34) exhibits reversal of effi-

cacy, appearing as an inverse agonist for cAMP production and a partial ago-

nist for ERK1/2. The ability of bPTH(3–34) and (D-Trp12,Tyr34)-bPTH

(7–34) to activate ERK1/2 in the absence of significant G protein-signaling

correlates with a high degree of sensitivity to arrestin2/3 knockdown,


suggesting that these ligands rely on arrestin coupling to activate ERK1/

2. Figure 13.6D shows the relationship between calcium signaling and

ERK1/2 activation. The two balanced ligands, hPTH(1–34) and

hPTH(1–31), appear similar, while Trp1-hPTHrp(1–36), bPTH(3–34),

and (D-Trp12,Tyr34)-bPTH(7–34) appear to be ERK1/2 biased, with par-

tial or full agonist activity in the ERK1/2 assay in the absence of a detectable

calcium response. hPTH(7–34), which has no significant activity in any of

the assays, still appears as a neutral antagonist.

3. DISCUSSION

The principal targets of PTH in vivo are kidney and bone, where its

actions promote a rise in serum calcium. In the kidney, PTH stimulates

reabsorption of filtered calcium by the cortical thick ascending limb of

the loop of Henle and distal convoluted tubule. In the proximal tubule, it

stimulates expression of the 1a-hydroxylase that converts 25(OH)-vitamin

D to its active form 1,25(OH)2-vitamin D, which in turn enhances intestinal

calcium absorption. The actions of PTH in bone are complex. PTH directly

stimulates osteoblasts, accelerating bone formation by increasing osteoblast

number and activity, promoting the deposition of new bone matrix and

accelerating the rate of mineralization (Dobnig & Turner, 1995; Schmidt,

Dobnig, & Turner, 1995). At the same time, PTH accelerates bone degra-

dation and the release of calcium locked in the mineralized skeleton by pro-

moting the recruitment, differentiation, and activity of bone-resorbing

osteoclasts. The effects of PTH on osteoclasts are indirect. Osteoclasts lack

PTH receptors, but respond to factors secreted by osteoblasts in response to

PTH, such as receptor activator of nuclear factor kB ligand. Because the ana-

bolic and catabolic effects of PTH are coupled, the net effect of PTH on

bone is dependent upon the kinetics of receptor activation, with intermit-

tent exposure leading to a net increase in bone formation, while continuous

exposure produces net bone loss and possible hypercalcemia (Dobnig &

Turner, 1995; Hock & Gera, 1992; Qin, Raggatt, & Partridge, 2004).

The skeletal effects of several conventional and biased PTH1R ligands

have already been determined in murine models, permitting some degree

of direct comparison between their in vitro efficacy profile and in vivo bio-

logical activity. PTH(1–34), which elicits the full range of PTH1R signaling

in vitro, also reproduces the full spectrum of PTH action in vivo. Mice given

daily injections of PTH(1–34) show increased indices of bone formation and

a net increase in bone mass (Dobnig & Turner, 1995). Osteoblast number,


osteoid surface, serum osteocalcin level, and mineral apposition rates, all

increase, as does trabecular bone volume and cortical thickness. At the same

time, indices of osteoclastic bone resorption rise, including osteoclast num-

ber and urinary deoxypyrodinoline level. Serum and urine calcium levels

also rise, reflecting the net effect of PTH(1–34) on bone resorption, intes-

tinal calcium absorption, and renal tubular calcium retention. PTH(1–31),

which in our screen appears to be a balanced partial agonist, also resembles

PTH(1–34) in vivo (Mohan et al., 2000). With several weeks treatment,

PTH(1–31) increases markers of bone formation as effectively as PTH

(1–34). The increase in bone resorption parameters is less dramatic

for PTH(1–31), yet the increase in bone density is also smaller. PTHrp

(1–36), which like Trp1-hPTHrp(1–36) is reportedly Gs pathway-selective,

is also anabolic in vivo, suggesting that Gq/11 signaling may be dispensable for

PTH1R actions in bone. In an ovariectomized rat model of bone loss, both

PTH(1–34) and PTHrp(1–36) increase indices of bone formation, bone

mass, and bone strength (Stewart et al., 2000). PTH(1–34), which produces

more sustained Gs-cAMP activation than PTHrp(1–36) in vitro, is more effi-

cacious in this model, where neither agent produces significant increases in

osteoclast activity.

UnlikeGq/11, thecapacity toactivateGs signalingappears tobe linked to the

anabolic effects of conventional PTH1R ligands. In mice, the N-terminal

truncated fragment, PTH(2–34),which is dramatically impaired inGs coupling

is far less efficacious than either PTH(1–34) or PTH(1–31) (Mohan et al.,

2000). Similarly, comparison of the ligand series, PTH(1–38), PTH(2–38),

and PTH(3–38) in rats further supports the conclusion that Gs signaling

is critical. Despite the capacity to activate PKC and simulate mitogenesis

in rat osteoblastic cells in vitro, PTH(3–38) produced no detectable anabolic

or catabolic effects on bone in vivo (Hilliker, Wergedal, Gruber, Bettica, &

Baylink, 1996).

However, results obtained with the putatively arrestin pathway-selective

biased agonist (D-Trp12, Tyr34)-bPTH(7–34) seem to contradict the general

conclusion that Gs activation is requisite for the anabolic actions of PTH.

Arrestin3 null mice exhibit higher basal rates of bone turnover and an

impaired anabolic response to PTH(1–34), with blunted increases in trabec-

ular bone volume and no change in cortical thickness compared to controls.

The attenuated response is associated with smaller changes in osteoblast

number and osteoid deposition, but preserved or exaggerated increases in

osteoclast number and urine deoxypyrodinoline (Bouxsein et al., 2005;

Ferrari et al., 2005; Gesty-Palmer et al., 2009). While this supports the


conclusion that PTH1R-mediated osteoclast activation is Gs-dependent, it

does not address whether the diminished anabolic PTH(1–34) response in

arrestin3 null mice reflects exaggerated cAMP signaling in the setting of

impaired arrestin-mediated desensitization or loss of arrestin-mediated signal-

ing. Paradoxically, intermittent administration of (D-Trp12, Tyr34)-bPTH

(7–34) to wild-type mice results in increased bone mass, characterized by

greater trabecular bone volume and increased osteoblast number, osteoid

surface, serum osteocalcin, and mineral apposition rate, with no significant

effect on osteoclast number or bone turnover markers (Gesty-Palmer et al.,

2009). All responses to (D-Trp12, Tyr34)-bPTH(7–34) are either absent or

reversed in arrestin 3 null mice, consistent with the hypothesis that arrestin sig-

naling in vivo contributes to the anabolic response to PTH, andwhen activated

in isolation is sufficient to promote osteoblastic bone formation but not to

stimulate osteoclastic bone resorption.

Results such as these, wherein two GPCR ligands with dramatically dif-

ferent in vitro activity profiles produce qualitatively similar biological responses

in vivo, suggest that there is something different about “arrestin-biased”

PTH1R activation at the tissue level that is not predicted by conventional

ligand screening. At present, it is unclear whether this means that changing

the balance of GPCR coupling is sufficient to produce markedly different

responses at the tissue level, where the strength/duration of signals originating

from proximal effectors are integrated to produce a response, or whether

biased ligands can couple GPCRs in “unnatural” ways, generating qualita-

tively different proximal efficacy. Either way, the finding that biasing GPCR

signaling can engender new/unexpected signaling outcomes has significant

implications for the rationale design of functionally selective drugs. While

it is clearly possible to tailor ligands to elicit specific efficacy profiles, we

are in many cases left with the quandary of not knowing which downstream

signals to favor and which to avoid. Thus, the greatest challenge at present is

not in detecting ligand bias, but in determining what efficacy profile is needed

to produce the optimal therapeutic response in any given setting.

ACKNOWLEDGMENTSThe authors thank Dr. Diane Gesty-Palmer (Duke University Medical Center) for helpful

advice and criticism and Allie Pinosky for technical assistance. The work was supported by

National Institutes of Health Grant R01 DK55524 (L. M. L.), the MUSC fluorescence

imaging plate reader (FLIPRTETRA) facility (S10 RR027777; L. M. L./T. A. M.), and the

Research Service of the Charleston, SC Veterans Affairs Medical Center (L. M. L./T. A. M.).

The contents of this chapter do not represent the views of the Department of Veterans Affairs

or the United States Government.


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