The human histamine H2-receptor couples more efficiently to Sf9 insect cell Gs-proteins than to...

The human histamine H2-receptor couples more efficiently to Sf9

insect cell Gs-proteins than to insect cell Gq-proteins: limitations

of Sf9 cells for the analysis of receptor/Gq-protein coupling

Christine Houston,* Katharina Wenzel-Seifert,* Tilmann Burckstummer and Roland Seifert*

*Department of Pharmacology and Toxicology, The University of Kansas, Lawrence, USA

Department of Molecular Biosciences, The University of Kansas, Lawrence, USA

Abstract

The human histamine H2-receptor (hH2R) couples to

Gs-proteins to activate adenylyl cyclase and to Gq-proteins to

activate phospholipase C, but phospholipase C activation has

not consistently been observed. The aim of this study was to

compare coupling of hH2R to insect and mammalian Gs- and

Gq-proteins in Spodoptera frugiperda (Sf9) cells. Interaction of

hH2R with mammalian G proteins was assessed with coex-

pressed proteins or receptor-Ga fusion proteins that enhance

coupling efficiency. hH2R efficiently coupled to insect

Gs-proteins to activate adenylyl cyclase. However, hH2R

poorly coupled to insect Gq-proteins as assessed by the lack

of enhancement of histamine-stimulated steady-state GTP

hydrolysis by regulators of G protein signaling (RGS proteins).

In contrast, RGS-proteins efficiently enhanced GTP hydrolysis

stimulated by the human platelet-activating factor receptor

(PAFR) and the histamine H1-receptor (H1R) from man and

guinea pig. The measurement of intracellular free Ca2+

concentration was not useful for studying receptor/Gq-protein

coupling. hH2R also efficiently interacted with mammalian

Gs-proteins, specifically with fused Gsa as assessed by

guanosine 5¢-O-(3-thiotriphosphate) (GTPcS)-sensitive high-

affinity agonist binding, agonist-stimulated [35S]GTPcS bind-

ing and adenylyl cyclase activation. In contrast, coupling of

hH2R to coexpressed and fused mammalian Gqa was poor.

However, our inability to reconstitute efficient coupling of

PAFR and H1R to mammalian Gqa indicated that a large

portion of the expressed G protein was functionally inactive.

Taken together, our data show that hH2R couples more effi-

ciently to insect cell Gs-proteins than to insect cell Gq-proteins.

Unfortunately, there are significant limitations in the useful-

ness of Sf9 cells for comparing the coupling of receptors to

mammalian Gs- and Gq-proteins and assessing Gq-mediated

activation of effector systems.

Keywords: fusion protein, Gq-protein, Gs-protein, H2-recep-

tor, regulator of G proteins, Sf9 insect cells.

J. Neurochem. (2002) 80, 678–696.

The histamine H2-receptor (H2R) is a prototypical neuro-

transmitter receptor that belongs to the superfamily of

G protein-coupled receptors (GPCRs) (Leurs et al. 1995; Hill

coupled receptor; gpH1R, guinea pig histamine H1-receptor; gpH2R,

guinea pig histamine H2-receptor; Gq-proteins, family of G proteins

(comprising mammalian Gqa, G11a, G14a, G15a and G16a and insect Gq-

proteins) that mediates phospholipase C activation; Gs-proteins, family

of G proteins (comprising mammalian Gsa-short, Gsa-long, Gsa-extralong and

Gaolf and insect Gs-proteins) that mediates adenylyl cyclase activation;

GsaL, long splice variant of the Gs-protein Gsa; GsaS, short splice variant

of the Gs-protein Gsa; GTPcS, guanosine 5¢-O-(3-thiotriphosphate);hH1R, human histamine H1-receptor; hH2R, human histamine H2-re-

ceptor; hH2R-GsaL, fusion protein containing the human histamine H2-

receptor and the long splice variant of Gsa; hH2R-GsaS, fusion protein

containing the human histamine H2-receptor and the short splice variant

of Gsa; HIS, histamine; HxR, non-specified histamine receptor;

[3H]MEP, [3H]mepyramine; MP, mastoparan (INLKALAALAKKIL);

PAF, platelet-activating factor; PAFR, human receptor for platelet-acti-

vating factor; PLC, phospholipase C; RGS4, regulator of G protein

signaling 4; rH2R, rat histamine H2-receptor; SDS–PAGE, sodium

dodecyl sulfate–polyacrylamide gel electrophoresis; TG, thapsigargin;

[3H]TIO, [3H]tiotidine.

Received September 7, 2001; revised manuscript received November 27,

2001; accepted November 29, 2001.

Address correspondence and reprint requests to Roland Seifert, De-

partment of Pharmacology and Toxicology, The University of Kansas,

Malott Hall, Room 5064, 1251 Wescoe Hall Drive, Lawrence, KS

66045–7582, USA. E-mail: [email protected]

Abbreviations used: AC, adenylyl cyclase; b2AR, human b2-adreno-

ceptor; b2AR-GsaL, fusion protein containing the human b2AR and the

long splice variant of Gsa; BSA, bovine serum albumin; [Ca2+]i, intra-

cellular free Ca2+ concentration; [3H]DHA, [3H]dihydroalprenolol; Ga,

non-specified G protein a-subunit; GAIP, G-alpha interacting protein, a

regulator of G protein signaling (RGS) protein; GPCR, G protein-

Journal of Neurochemistry, 2002, 80, 678–696

678 Ó 2002 International Society for Neurochemistry, Journal of Neurochemistry, 80, 678–696

et al. 1997). The H2R is widely distributed in the brain, with

the highest densities found in the basal ganglia, hippocampus,

amygdala and cortex (Traiffort et al. 1992a). There is evidence

to support the assumption that the human H2R (hH2R) plays a

role in the development of Huntington’s chorea (Martinez-Mir

et al. 1993), autism (Linday et al. 2001), schizophrenia

(Martinez 1999) and brain tumors (Panula et al. 2000), but

overall, the function of the hH2R in the brain is still poorly

understood. In order to better understand the function of the

hH2R in brain functions it is essential to have detailed

knowledge on the signaling pathways regulated by this

GPCR. It has been consistently reported that the hH2R

activates adenylyl cyclase (AC) through the G protein Gs both

in native and recombinant systems (Bristow et al. 1982;

Gespach and Abita 1982; Mitsuhashi et al. 1989; Leurs et al.

1994; Alewijnse et al. 1998; Wang et al. 2000). In addition to

AC activation, the hH2R can also mediate phospholipase C

(PLC) activation via Gq-proteins, but PLC activation has not

consistently been observed (Mitsuhashi et al. 1989; Seifert

et al. 1992; Leurs et al. 1994; Burde and Seifert 1996; Wang

et al. 2000). Moreover, the functional importance of hH2R-

mediated PLC activation is elusive.

The uncertainty regarding Gq-coupling of the hH2R

prompted us to compare Gs- and Gq-coupling of the hH2R

under defined experimental conditions. We used Sf9 insect

cells as an expression system and studied H2R/G protein

coupling in three different settings. First, we assessed

coupling of hH2R to the endogenous insect cell G proteins.

Second, we analyzed coupling of hH2R to coexpressed

mammalian G proteins. Third, we studied coupling of hH2R

fused to mammalian Ga-subunits. GPCR-Ga fusion proteins

ensure close physical proximity between GPCR and Ga and

ensure efficient interaction (Seifert et al. 1999; Milligan

2000). While efficient coupling in GPCR-Gsa fusion proteins

is well established (Bertin et al. 1994; Seifert et al. 1998b;

Unson et al. 2000; Wenzel-Seifert and Seifert 2000; Liu et al.

2001b), efficient coupling in GPCR-Gq/11a fusion proteins has

only recently been documented (Holst et al. 2001; Stevens

et al. 2001). As read-outs for GPCR/G protein coupling, we

determined GTPcS-sensitive high-affinity agonist binding as

well as agonist-stimulated [35S]GTPcS binding, steady-state

GTP hydrolysis and effector system activation. Here, we show

that the hH2R couples more efficiently to insect cell Gs-pro-

teins than to insect cell Gq-proteins, but there are limitations in

the usefulness of Sf9 cells as a model system for comparing

the coupling of GPCRs to mammalian Gs- and Gq-proteins

and assessing Gq-mediated activation of effector systems.

Materials and methods

Materials

The cDNA for hH2R was kindly provided by Dr I. Gantz

(University of Michigan Medical School and Ann Arbor VA

Medical Center, Ann Arbor, MI, USA) (Gantz et al. 1991). The

cDNAs for hH1R and gpH1R were kindly provided by Dr H. Fukui

(Department of Pharmacology, University of Tokushima, Tokushi-

ma, Japan) (Horio et al. 1993; Fukui et al. 1994). Baculoviruses

encoding mammalian Gqa, RGS4 and GAIP were kindly donated by

Dr E. Ross (Department of Pharmacology, University of South-

western Medical Center, Dallas, TX, USA). The baculoviruses

encoding GsaS and Gia2 were kindly provided by Drs R. Sunahara

and A. G. Gilman (Department of Pharmacology, University of

Southwestern Medical Center, Dallas, TX, USA). The baculovirus

encoding G protein subunits b1c2 was a gift from Dr P. Gierschik

(Department of Pharmacology and Toxicology, University of Ulm,

Germany). The baculovirus encoding rH2R was kindly donated by

Drs C. Harteneck and G. Schultz (Department of Pharmacology,

Free University of Berlin, Germany). The construction of FLAG

epitope- and hexahistidine-tagged cDNAs for b2AR-GsaL, hH2R-

GsaS, hH2R, gpH2R and PAFR was described elsewhere (Seifert

et al. 1998a; Kelley et al. 2001; Seifert and Wenzel-Seifert 2001).

Fura-2-acetoxymethylester was from Molecular Probes (Eugene,

OR, USA). The anti-FLAG Ig (M1 monoclonal antibody), HIS, TG,

MP, GdCl3, LaCl3 and PAF were from Sigma (St. Louis, MO, USA).

Octopamine was from RBI (Natick, MA, USA). The anti-Gsa Ig

(C-terminal) and the anti-Gqa Igs 371730, 371752 and 371754 and

recombinant mammalian Gqa purified from E. coli. were from

Calbiochem (La Jolla, CA, USA). The K20 Ig, anti-RGS4 Ig

(N-terminal) and anti-GAIP Ig (N-terminal) were from Santa Cruz

(Santa Cruz, CA, USA). [a-32P]ATP (3000 Ci/mmol), [35S]GTPcS(1100 Ci/mmol), [c-32P]GTP (6000 Ci/mmol), [3H]DHA

(85–90 Ci/mmol) and [3H]TIO (90 Ci/mmol) were from Perkin

Elmer Life Sciences (Boston, MA, USA). [3H]MEP (30 Ci/mmol)

was from Amersham Pharmacia Biotech (Piscataway, NJ, USA). All

unlabeled nucleotides were of the highest purity available and were

either from Roche (Indianapolis, IN, USA) or Sigma. All restriction

enzymes and T4 DNA ligase were from New England Biolabs

(Beverly, MA, USA). Cloned Pfu DNA polymerase was from

Stratagene (La Jolla, CA, USA). The GC-RICH PCR-System was

from Roche. Unless specifically stated otherwise, PCRs were

conducted with Pfu DNA polymerase. Sf9 cells were from the

American Type Cell Culture Collection (Rockville, MD, USA). All

other reagents were of the highest purity available and were from

Fisher (Pittsburgh, PA, USA) or Sigma.

Construction of FLAG epitope- and hexahistidine-tagged

cDNA for hH2R-Gqa

In PCR 1, the C-terminal portion of hH2R was amplified using

pGEM-3Z-SF-hH2R as template, a sense primer annealing 5¢ of theengineered EcoR V site of the hH2R and an antisense primer

annealing with the hexahistidine tag. In PCR 2, the sequence of Gqa

was amplified, using pGEM-3Z-SF-b2AR-Gqa as template, a sense

primer annealing with the hexahistidine tag and an antisense primer

annealing with the 5 C-terminal amino acids of Gqa, the stop codon

and a BamH I site. In PCR 3, the products of PCRs 1 and 2 annealed

in the hexahistidine region, and the sense primer of PCR 1 and the

antisense primer of PCR 3 were used. In this way, a fragment

encoding the C-terminal portion of the hH2R, a hexahistidine tag,

Gqa, a stop codon and a BamH I site was created. This fragment was

digested with EcoR V and BamH I and cloned into pGEM-3Z-

SFhH2R digested with EcoR V and BamH I. In this way, the

G protein coupling of human H2-receptor 679

Ó 2002 International Society for Neurochemistry, Journal of Neurochemistry, 80, 678–696

full-length cDNA for hH2R-Gqa was created. pGEM-3Z-SF-hH2R-

Gqa was digested with Nco I and BamH I to recover the fusion

protein cDNA and cloned into the baculovirus transfer vector pVL

1392-SF-b2AR-Gia2 digested with Nco I and BamH I. PCR-

generated DNA sequences were confirmed by extensive restriction

enzyme analysis and enzymatic sequencing.

Construction of FLAG epitope- and hexahistidine-tagged

cDNAs for hH1R and gpH1R

A DNA sequence encoding the cleavable signal peptide from

influenza hemagglutinin (S) followed by the FLAG epitope (F),

which is recognized by the M1 monoclonal antibody, was placed 5¢of the start codon of the hH1R and gpH1R in order to enhance GPCR

expression and allow immunological detection. We also added a

hexahistidine tag to the C-terminus of hH2R to allow future

purification and to provide additional protection against proteolysis.

The GPCR modifications were generated by sequential overlap-

extension PCRs.

In PCR 1A, the DNA for the SF region was amplified with

pGEM3Z-SF-hH2R-Gsas as template by using a sense primer 5¢ ofthe Sac I site near the SF sequence and an antisense primer

encoding the last 19 bp of the SF. In PCR 1B, the DNA sequence

of the hH1R was amplified using pDKCR-hH1R as template. The

sense primer annealed with the first 17 bp of the 5¢-end of the hH1R

and included the last 15 bp of the SF in its 5¢-extension. The

antisense primer encoded the five C-terminal amino acids of the

hH1R, a hexahistidine tag, the stop codon and an Xba I site. The

PCR was conducted with the GC-RICH PCR-System using 1 mM

GC-RICH resolution solution, denaturation at 95°C for 45 s,

annealing at 50°C for 45 s and extension at 72°C for 90 s for a total

of 30 cycles. In PCR 2, the products of PCR 1A and 1B were used

as templates and annealed in the F region. Amplification was

initiated with the sense primer of PCR 1 A and the antisense primer

of PCR 1B. In this way, a fragment encoding the signal sequence,

the FLAG epitope and hH1R cDNA with a hexahistidine tag

followed by an Xba I site was obtained. This fragment was digested

with Hind III and Xba I and cloned into pGEM-3Z-SF-human

formyl peptide receptor-6His digested with Hind III and Xba I.

pGEM-3Z-SF-hH1R was digested with Hind III and Xba I to

recover the hH1R cDNA and cloned into the baculovirus transfer

vector pVL 1392-SF-b2AR-Gia2 digested with Sac I and Xba I.

Since the N-terminal restriction site of the hH1R cDNA (Hind III)

did not match the corresponding restriction site in the transfer

vector (Sac I), a blunt-end ligation was performed. PCR-generated

DNA sequences were confirmed by extensive restriction enzyme

analysis and enzymatic sequencing.

In PCR 3, the DNA sequence of the gpH1R was amplified using

pEFBOS-gpH1R as template. The sense primer annealed with the

first 17 bp of the 5¢-end of the gpH1R and included the last 15 bp of

the SF in its 5¢-extension. The antisense primer encoded the five

C-terminal amino acids of the gpH1R, a hexahistidine tag, the stop

codon and an Xba I site. The PCR was conducted using the

GC-RICH PCR-System with 1 mM GC-RICH resolution solution,

denaturation at 95°C for 45 s, annealing at 50°C for 45 s and

extension at 72°C for 90 s for a total of 30 cycles.

In PCR 4, the sense primer of PCR 1 A and the antisense

primer of PCR 3 were used. The PCR was conducted using the

GC-RICH PCR-System with 1 mM GC-RICH resolution solution,

denaturation at 95°C for 45 s, annealing at 50°C for 45 s and

extension at 72°C for 90 s for a total of 30 cycles. In this way, a

fragment encoding the signal sequence, the FLAG epitope and

gpH1R cDNA with a hexahistidine tag followed by an Xba I site

was obtained. This fragment was digested with Sac I and Xba I

and cloned into pGEM-3Z-SF-human formyl peptide receptor-

6His digested with Sac I and Xba I. In this way, the full-length

cDNA for gpH1R was created. pGEM-3Z-SF-gpH1R was digested

with Sac I and Xba I to recover the gpH1R cDNA and cloned into

the baculovirus transfer vector pVL 1392-SF-b2AR-Gia2 digested

with Sac I and Xba I. PCR-generated DNA sequences were

confirmed by extensive restriction enzyme analysis and enzymatic

sequencing.

Generation of recombinant baculoviruses, cell culture

and membrane preparation

Sf9 cells were cultured in 250 mL disposable Erlenmeyer flasks at

28°C under rotation at 125 r.p.m. in SF 900 II medium (Life

Technologies, Carlsbad, CA, USA) supplemented with 5% (v/v)

fetal calf serum (Bio Whittaker, Walkersville, MD, USA) and

0.1 mg/mL gentamicin. Cells were maintained at a density of

0.5–6.0 · 106 cells/mL. Recombinant baculoviruses were generated

in Sf9 cells using the BaculoGOLD transfection kit (Pharmingen,

San Diego, CA, USA) according to the manufacturer’s instructions.

After initial transfection, high-titer virus stocks were generated by

two sequential virus amplifications. In the first amplification, cells

were seeded at 2.0 · 106 cells/mL and infected with a 1 : 100

dilution of the supernatant from the initial transfection. Cells were

cultured for 7 days, resulting in the death of virtually the entire cell

population. The supernatant fluid of this infection was harvested and

stored under light protection at 4°C. In a second amplification, cells

were seeded at 3.0 · 106 cells/mL and infected with a 1 : 20

dilution of the supernatant from the first amplification. Cells were

cultured for 48 h, and the supernatant fluid was harvested. After the

48 h culture, the majority of cells showed signs of infections (e.g.

altered morphology, viral inclusion bodies), but most of the cells

were still intact. The supernatant fluid from the second amplification

was also stored under light protection at 4°C and was the routine

virus stock for membrane preparations and intact cell experi-

ments. For infection, cells were sedimented by centrifugation and

suspended in fresh medium at 3.0 · 106 cells/mL. For membrane

preparation, cells were infected with a 1 : 100 dilution of high-titer

baculovirus stocks encoding GPCRs, Ga, b1c2, RGS proteins and/or

GPCR-Ga fusion proteins. Cells were cultured for 48 h before

membrane preparation. Sf9 membranes were prepared as described

(Seifert et al. 1998a), using 1 mM EDTA, 0.2 mM phenyl-

methylsulfonyl fluoride, 10 lg/mL benzamidine and 10 lg/mL

leupeptin as protease inhibitors. Membranes were suspended in

binding buffer (12.5 mM MgCl2, 1 mM EDTA and 75 mM Tris/HCl,

pH 7.4) and stored at ) 80°C until use. For determination of [Ca2+]i,

cells were infected with baculoviruses for 18–48 h and virus

dilutions of 1 : 100–1 : 10 000.

Receptor ligand binding assays

Membranes were thawed and sedimented by a 15-min centrifugation

at 4°C and 15 000 · g to remove residual endogenous guanine

nucleotides as far as possible. Membranes were resuspended in

binding buffer. [3H]TIO binding was carried out as described

680 C. Houston et al.


(Kelley et al. 2001; Wenzel-Seifert et al. 2001). Briefly, each tube

(total volume 250 lL) contained 200–250 lg of protein. Incuba-

tions were conducted for 90 min at 25°C and shaking at 250 r.p.m.

For H2R saturation binding experiments, tubes contained 1–20 nM

[3H]TIO plus unlabeled TIO to obtain final ligand concentrations of

up to 300 nM. Non-specific [3H]TIO binding was determined in the

presence of 100 lM unlabeled TIO and amounted to �55–65% of

total binding with high [3H]TIO concentrations (Kelley et al. 2001;

Wenzel-Seifert et al. 2001). In agonist competition binding experi-

ments, reaction mixtures contained 10 nM [3H]TIO and unlabeled

HIS at various concentrations without or with GTPcS (10 lM).

Bound [3H]TIO was separated from free [3H]TIO by filtration

through GF/C filters, followed by three washes with 2 mL of

binding buffer (4°C). Filter-bound radioactivity was determined by

liquid scintillation counting. The experimental conditions chosen

ensured that not more than 5% of the total amount of [3H]TIO added

to binding tubes bound to filters. Non-specific binding with 10 nM

[3H]TIO amounted to �10–15% of total [3H]TIO binding.

In [3H]MEP binding assays each tube (total volume 500 lL)contained 20–25 lg of protein. Incubations were conducted for

90 min at 25°C and shaking at 250 r.p.m. For H1R saturation

binding experiments, tubes contained 0.2–20 nM [3H]MEP.

Non-specific binding was determined in the presence of 10 lM

unlabeled MEP and amounted to �50% of total binding with high

[3H]MEP concentrations. In agonist competition binding experi-

ments, reaction mixtures contained 2 nM [3H]MEP and unlabeled

HIS at various concentrations without or with GTPcS (10 lM).

Bound [3H]MEP was separated from free [3H]MEP by filtration

through GF/C filters, followed by three washes with 2 mL of

binding buffer (4°C). Filter-bound radioactivity was determined by

liquid scintillation counting. The experimental conditions chosen

ensured that not more than 5–10% of the total amount of [3H]MEP

added to binding tubes bound to filters. Non-specific binding with

2 nM [3H]MEP amounted to �10% of total [3H]MEP binding.

The expression level of b2AR and b2AR-GsaL was determined

with 10 nM [3H]DHA as radioligand as described (Seifert et al.

1998a).

Steady-state GTPase activity assay

GTPase activity was determined as described (Kelley et al. 2001;

Liu et al. 2001b). Briefly, membranes were thawed, sedimented and

resuspended in 10 mM Tris/HCl, pH 7.4. Assay tubes contained Sf9

membranes (10 lg of protein/tube) expressing various GPCRs

without or with RGS proteins and/or mammalian G proteins,

1.0 mM MgCl2, 0.1 mM EDTA, 0.1 mM ATP, 100 nM GTP, 1 mM

adenylyl imidodiphosphate, 5 mM creatine phosphate, 40 lg of

creatine kinase and 0.2% (w/v) bovine serum albumin (BSA) in

50 mM Tris/HCl, pH 7.4. Reactions mixtures additionally contained

solvent (basal) or the appropriate agonists at a maximally effective

concentration. Reaction mixtures (80 lL) were incubated for 3 min

at 25°C before the addition of 20 lL of [c-32P]GTP (0.2–0.5

lCi/tube). All stock- and work dilutions of [c-32P]GTP were

prepared in 20 mM Tris/HCl, pH 7.4. Reactions were conducted for

20 min at 25°C. Reactions were terminated by the addition of

900 lL of a slurry consisting of 5% (w/v) activated charcoal and

50 mM NaH2PO4, pH 2.0. Charcoal absorbs nucleotides but not Pi.

Charcoal-quenched reaction mixtures were centrifuged for 15 min at

room temperature at 15 000 g. Supernatant fluid of the reaction

mixtures was removed (700 lL), and 32Pi was determined by liquid

scintillation counting. Enzyme activities were corrected for sponta-

neous degradation of [c-32P]GTP. Spontaneous [c-32P]GTP degra-

dation was determined in tubes containing all of the above described

components plus a very high concentration of unlabeled GTP

(1 mM) that, by competition with [c-32P]GTP, prevents [c-32P]GTPhydrolysis by enzymatic activities present in Sf9 membranes.

Spontaneous [c-32P]GTP degradation was < 1% of the total amount

of radioactivity added using 20 mM Tris/HCl, pH 7.4, as solvent for

[c-32P]GTP. The experimental conditions chosen ensured that not

more than 10% of the total amount of [c-32P]GTP added was

converted to 32Pi.

[35S]GTPcS binding assay

[35S]GTPcS binding was determined as described (Seifert et al.

1998a; Wenzel-Seifert et al. 2001). Briefly, membranes were thawed

and sedimented by a 15-min centrifugation at 4°C and 15 000 · g

to remove residual endogenous guanine nucleotides as far as

possible and resuspended in binding buffer. Reaction mixtures (total

volume 500 lL) contained Sf9 membranes (15–20 lg of protein/

tube) expressing various proteins in binding buffer supplemented

with 0.05% (w/v) BSA, 0.4 nM [35S]GTPcS and unlabeled GDP at

various concentrations. Reaction mixtures additionally contained

distilled water (basal) or HIS at a saturating concentration (100 lM).

Incubations were conducted for 60 min at 25°C and shaking at

250 r.p.m. Bound [35S]GTPcS was separated from free [35S]GTPcSby filtration through GF/C filters, followed by three washes with

2 mL of binding buffer (4°C). Filter-bound radioactivity was

determined by liquid scintillation counting. The experimental

conditions chosen ensured that no more than 10% of the total

amount of [35S]GTPcS added was bound to filters. Non-specific

binding with 0.4 nM [35S]GTPcS amounted to < 0.1% of total

[35S]GTPcS binding. Bound [35S]GTPcS was separated from free

[35S]GTPcS by filtration through GF/C filters, followed by three

washes with 2 mL of binding buffer (4°C). Filter-bound radio-

activity was determined by liquid scintillation counting.

In time course experiments, Sf9 membranes expressing various

proteins were suspended in 1500 lL of binding buffer supple-

mented with 1 nM [35S]GTPcS plus 9 nM unlabeled GTPcS, 1 lM

GDP, and distilled water (basal) or HIS (100 lM). Aliquots of

200 lL containing 25 lg of protein were withdrawn at different

time points.

AC activity assay

AC activity in Sf9 membranes was determined as described

(Alvarez and Daniels 1990; Liu et al. 2001b). Briefly, membranes

were thawed and sedimented by a 15-min centrifugation at 4°C and

15 000 g to remove residual endogenous guanine nucleotides as far

as possible and resuspended in binding buffer. Tubes contained Sf 9

membranes (30–50 lg of protein/tube) expressing various proteins,

5 mM MgCl2, 0.4 mM EDTA and 30 mM Tris/HCl, pH 7.4. Assay

tubes containing membranes and various additions in a total volume

of 30 lL were incubated for 3 min at 37°C before starting reactions

by the addition of 20 lL of reaction mixture containing (final)

[a-32P]ATP (1.0–1.5 lCi/tube) plus 40 lM unlabeled ATP, 2.7 mM

mono(cyclohexyl)ammonium phosphoenolpyruvate, 0.125 IU of

pyruvate kinase, 1 IU of myokinase and 0.1 mM cAMP. Reactions

were conducted for 20 min at 37°C. Reactions were terminated by



the addition of 20 lL of 2.2 N HCl. Denatured protein was

sedimented by a 3-min centrifugation at 25°C and 15 000 · g.

Sixty-five microliters of the supernatant fluid were applied onto

disposable columns filled with 1.3 g of neutral alumina (Sigma

A-1522, super I, WN-6). [32P]cAMP was separated from

[a-32P]ATP by elution of [32P]cAMP with 4 mL of 0.1 M

ammonium acetate, pH 7.0 (Alvarez and Daniels 1990). Recovery

of [32P]cAMP was �80%. Blank values were routinely �0.01% of

the total amount of [a-32P]ATP added. [32P]cAMP was determined

by liquid scintillation counting. The experimental conditions chosen

ensured that not more than 1–3% of the total amount of [a-32P]ATPadded was converted to [32P]cAMP.

Determination of [Ca2+]i

Sf9 cells were centrifuged for 5 min at 500 · g at 20°C. After thecentrifugation, the supernatant fluid was discarded, and cells were

suspended at 2.0 · 106 cells/mL in MBS buffer [10 mM NaCl,

60 mM KCl, 17 mM MgCl2, 4 mM D-glucose, 110 mM sucrose,

10 mM MES and 0.1% (w/v) BSA, pH 6.2] supplemented with

1 mM CaCl2 and 2 lM fura-2-acetoxymethylester. Cells were

incubated for 45 min at 25°C under light protection. After loading

with fura-2, the same volume of MBS buffer as used for dye loading

was added to cells, followed by centrifugation for 5 min at 500 g

at 20°C. The supernatant fluid was discarded, and cells were

suspended in MBS buffer at a density of 2.0 · 106 cells/mL. Cells

(2.0 · 106) were transferred into microcentrifuge tubes and centri-

fuged for 5 min at 1000 g at 20°C. The supernatant fluid was

removed, and cells were stored at 4°C under light protection until

use. Cells were kept at 4°C for up to 2 h. Immediately prior to use

each cell pellet was suspended in 1 mL of MBS buffer and

transferred into a UV cuvette. An additional 960 lL of MBS buffer

supplemented with (final concentration) 1 mM CaCl2 or 1 mM

EGTA were added to the cuvette. In some experiments, GdCl3 or

LaCl3 (10 lM each) were added to cuvettes at this point. Cuvettes

were placed into a Shimadzu RF-5000 spectrofluorometer, and

fluorescence (kex ¼ 340 nm and kem ¼ 500 nm) was continuously

recorded at 22°C under constant stirring of cells at 1000 r.p.m. After

a 1-min recording of basal [Ca2+]i, stimulus was added to cells, and

fluorescence was recorded for an additional 3–4 min Thereafter,

Fmax was determined by adding Triton X-100 (0.1%, w/v) to cells.

Fmin was determined by adding EGTA (40 mM, pH 8.0) to cells.

[Ca2+]i-values were calculated as described (Seifert et al. 1992)

using a Kd value of 278 nM for binding of Ca2+ to fura-2 (Harteneck

et al. 1995).

Sodium dodecyl sulfate–polyacrylamide gel electrophoresis

(SDS–PAGE) and immunoblot analysis

Membrane proteins were separated on SDS–polyacrylamide gels

containing 10% (w/v) acrylamide. Proteins were then transferred

onto Immobilon P transfer membranes (Millipore, Bedford, MA,

USA). Membranes were reacted with M1 antibody, anti-Ga Igs

(from 1 : 100 to 1 : 1000) or anti-RGS protein Igs (1 : 100).

Immunoreactive bands were visualized by sheep anti-mouse IgG

(M1 antibody), donkey anti-rabbit IgG (anti-Ga Igs) and donkey

anti-goat IgG (anti-RGS protein Igs), respectively, coupled to

peroxidase, using o-dianisidine and H2O2 as substrates. Secondary

antibodies were used at 1 : 500–1 : 1000 dilutions.

Miscellaneous

Protein concentrations were determined using the Bio-Rad DC

protein assay kit (Bio-Rad, Hercules, CA, USA). The analysis of

hH2R-Gqa expression by reverse transcription (RT)-PCR and

digestion with restriction enzymes was performed as described

previously for the b2AR-Gqa fusion protein (Wenzel-Seifert and

Seifert 2000). Data shown in Figs 4 and 5 and Table 1 were

analyzed by non-linear regression with the Prism III program

(GraphPad, Prism, San Diego, CA, USA).

Results and discussion

Detection of recombinant proteins in Sf9 cell membranes

by radioligand binding and immunoblotting

In membranes expressing hH2R, the antagonist radioligand

[3H]TIO bound with a Kd value of 63.0 ± 9.3 nM and a

Bmax of 0.62 ± 0.04 pmol/mg (mean ± SD, n ¼ 3). The

corresponding values for hH2R-GsaS were 32.0 ± 4.6 nM

and 0.43 ± 0.02 pmol/mg (Wenzel-Seifert et al. 2001),

and for hH2R-Gqa the values were 36.7 ± 4.9 nM and

Table 1 Non-linear regression analysis of the binding properties of HIS in Sf9 membranes expressing hH2R without or with various G protein

subunits and in membranes expressing hH2R-Ga fusion proteins

Construct Kh (lM) Kl (lM) Rh (%) KhGTPcS (lM) KlGTPcS (lM) RhGTPcS (%)

hH2R – 1.09 (0.82–1.46) – – 1.44 (1.07–1.93) –

hH2R + GsaS 0.09 (0.02–0.58) 4.60 (1.41–15.0) 38.8 (12.8–64.9) – 1.78 (1.17–2.69) –

hH2R-GsaS 0.08 (0.05–0.13) 8.26 (4.00–17.0) 62.4 (54.3–70.6) 0.04 (0.01–0.40) 5.84 (3.67–9.30) 22.2 (11.1–33.3)

hH2R + Gqa + b1c2 – 1.80 (01.00–3.24) – – 2.34 (1.03–5.28) –

hH2R-Gqa + b1c2 – 1.24 (0.89–1.71) – – 2.06 (1.51–2.81) –

Agonist competition binding was determined as described in Materials and methods. The data shown in Fig. 4 were analyzed by non-linear

regression for best fit to monophasic or biphasic competition isotherms (F-test). Data shown are the means from three to seven independent

experiments performed in duplicates. Numbers in parentheses represent the 95% confidence intervals. Kh and Kl designate the dissociation

constants for the high- and low-affinity state of hH2R constructs, respectively. %Rh indicates the percentage of high-affinity binding sites. The

corresponding values obtained in the presence of GTPcS (10 lM) are referred to as KhGTPcS, KlGTPcS and percentage RhGTPcS, respectively. If data

were best fit to monophasic competition curves, data are listed under Kl and KlGTPcS, respectively.



0.17 ± 0.05 pmol/mg. The expression levels of hH2R con-

structs in the [3H]TIO saturation binding assay, specifically

those of fusion proteins, were lower than the expression

levels estimated by immunoblot analysis (hH2R,�1 pmol/mg,

Fig. 1a; hH2R-GsaS, �2 pmol/mg, Fig. 1b; hH2R-Gqa,

�1 pmol/mg, Fig. 1c). Similar observations were made

before for other H2R-Ga fusion proteins (Kelley et al.

2001; Wenzel-Seifert et al. 2001). These data show that

fusion of H2R to Ga diminishes accessibility of H2R for

[3H]TIO. The molecular basis for the reduced labeling of

hH2R-Ga fusion protein molecules by [3H]TIO relative to

non-fused hH2R is unknown, but from a practical standpoint

of view, these findings render the [3H]TIO saturation binding

assay less feasible for assessing the expression hH2R

constructs than the immunoblot analysis.

In the immunoblot, the expression of hH2R constructs was

estimated using b2AR or b2AR-GsaL expressed at defined

levels (determined by [3H]DHA saturation binding) as

standard. To estimate Gqa expression levels, we used

recombinant Gqa purified from E. coli as standard because

anti-Gqa Igs are not sensitive enough to detect GPCR-Gqa

fusion proteins (Wenzel-Seifert and Seifert 2000). We

estimated the expression levels of proteins by visual

inspection of gels because the varying shapes of proteins

caused by dimerization or different glycosylation rendered

densitometric analysis ambiguous.

First, we performed immunoblots with the M1 monoclonal

antibody that recognizes the N-terminal FLAG epitope

attached to fused and non-fused GPCRs. As reported before

(Seifert et al. 1998a), the b2AR migrates as a doublet of

�52 kDa, representing differently glycosylated forms of the

GPCR (Fig. 1a). Similarly, hH2R migrated as a doublet of

�35 kDa. In membranes expressing hH2R we also detected a

doublet in the �60-kDa region. Most likely, those bands

represent hH2R dimers. Dimers that are resistant to denatur-

ation by SDS and reducing agents were also observed for the

canine H2R (Fukushima et al. 1997). The b2AR used as

standard in Fig. 1a was expressed at a level of�3.9 pmol/mg.

Using identical amounts of protein loaded, we estimated that

hH2R (monomer plus dimer) was expressed at a level of

�1 pmol/mg. This value is �40% lower than the value

obtained in the [3H]TIO saturation binding experiments (see

above). Similar expression levels of hH2R were obtained

when the GPCR was coexpressed with GsaS and Gqa (data

not shown).

The apparent molecular masses of GsaS and GsaL are

�45 kDa and 52 kDa, respectively (Graziano et al. 1989).

The molecular mass of b2AR-GsaL was �104 kDa, and the

molecular mass of hH2R-GsaS was �76 kDa (Fig. 1b). It

should be noted that no bands below the fusion proteins were

Fig. 1 Analysis of the expression of hH2R, hH2R-GsaS and hH2R-Gqa

in Sf9 membranes. Sf9 membranes expressing various proteins were

prepared, separated by SDS–PAGE on gels containing 10% (w/v)

acrylamide, transferred onto Immobilon P membranes and probed with

the anti-FLAG Ig (M1 antibody). Shown are the horseradish peroxi-

dase-reacted membranes. Numbers on the left of membranes desig-

nate masses of marker proteins in kDa. The amounts of protein loaded

onto each lane are given below membranes. In (a), b2AR expressed at

a level of 3.9 pmol/mg was used as standard. In (b), b2AR-GsaL

expressed at a level of 8.6 pmol/mg was used as standard. In (c),

b2AR expressed at 12.1 pmol/mg was used as standard.



detected, indicating that fusion proteins were not degraded.

b2AR-GsaL used as standard in Fig. 1(b) was expressed at a

level of �8.6 pmol/mg. Comparing identical amounts of

protein loaded, we estimated that hH2R-GsaS was expressed

at a level of �2 pmol/mg. The [3H]TIO saturation binding

experiments gave a �5-fold lower expression level of hH2R-

GsaS (see above).

The molecular mass of Gqa is �42 kDa (Hepler et al.

1993). As predicted, the molecular mass of hH2R-Gqa was

�77 kDa, and no degradation products were detected

(Fig. 1c). Using b2AR expressed at 12.1 pmol/mg as stan-

dard and loaded at a four times lower amount than hH2R-

Gqa, the expression level of hH2R-Gqa was estimated to be

� 1 pmol/mg. The [3H]TIO saturation binding experiments

gave a �six-fold lower expression level of hH2R-Gqa (see

above).

As expected (Graziano et al. 1989), the anti-Gsa Ig

(recognizing the C-terminal sequence RMHLRQYELL)

detected a �45-kDa antigen in membranes coexpressing

hH2R and GsaS (Fig. 2a). In a previous study we reported

that GsaL is expressed at high levels (�100–150 pmol/mg)

in Sf9 membranes (Seifert et al. 1998a). GsaS was

expressed at much lower levels in Sf9 membranes

(�1 pmol/mg) than GsaL using b2AR-GsaL expressed at

7.0 pmol/mg as standard. The reason for the low expression

level of GsaS is not known, but we also observed that the

maximum expression levels of b2AR-GsaS are lower than

the maximum expression levels of b2AR-GsaL (Seifert et al.

1998b). However, as will be described below, despite the

low expression of GsaS, coupling of hH2R to this G protein

was quite efficient.

hH2R-GsaS and hH2R-GsaL were also detected by anti-Gsa

Ig, and as expected (Graziano et al. 1989; Seifert et al.

1998b), hH2R-GsaL migrated slightly slower in SDS–PAGE

than hH2R-GsaS (Fig. 2b). Expected, too, was the finding that

hH2R-GsaL was expressed at higher levels than hH2R-GsaS.

In a recent study we reported that in contrast to the

corresponding b2AR-Gsa fusion proteins (Seifert et al.

1998b; Wenzel-Seifert and Seifert 2000), there are only

few functional differences between hH2R-GsaS and hH2R-

GsaL (Wenzel-Seifert et al. 2001). Therefore, in the present

study, we focused on the analysis of the coupling of hH2R to

GsaS and did not include additional data on hH2R-GsaL. As

reported before (Seifert et al. 1998a), the anti-Gsa Ig

(C-terminal) did not react with the endogenous Gs-protein of

the insect cells (Fig. 2c). The antibody K20, reacting with the

internalsequence85–103ofGsaS(KEAIETIVAAMSNLVPPVE)

(Krieger-Brauer et al. 1999) reacted with GsaS but failed to

recognize insect Gsa, too (Fig. 2d).

We also studied the expression of Gq-proteins in Sf9 cells.

Recombinant mammalian Gqa purified from E. coli migrated

as a �49-kDa protein in SDS–PAGE and reacted with

antibody 371730, raised against recombinant mammalian

Gqa (Fig. 3a), antibody 371752, recognizing the internal

sequence (115–133) EVDVEKVSAFENPYVDAIK (Fig. 3b)

and antibody 371754, recognizing the internal sequence

(283–300) IMYSHLVDYFPEYDGPQR (Fig. 3c). However,

Fig. 2 Analysis of the expression of GsaS and hH2R-GsaS in Sf9

membranes. Sf9 membranes expressing various proteins were pre-

pared, separated by SDS–PAGE on gels containing 10% (w/v)


the anti-Gsa Ig (C-terminal) (a–c) or antibody K20 (d). Shown are the

horseradish peroxidase-reacted membranes. Numbers on the left of

membranes designate masses of marker proteins in kDa. In (a), the

amounts of protein loaded onto each lane are given below the mem-

brane. In (b–d), 100 lg of protein were loaded onto each lane. In (a),

b2AR-GsaL expressed at a level of 7.0 pmol/mg was used as standard.



all anti-Gqa Igs studied failed to detect the endogenous Gq-

protein of Sf9 cells. In contrast, all anti-Gqa Igs studied (data

only shown for antibody 371754) detected mammalian Gqa

expressed in Sf9 cells that migrated as a �43-kDa protein

(Fig. 3d). The electrophoretic mobility of mammalian Gqa in

Sf9 membranes is in accordance with previous results

(Hepler et al. 1993). The difference in migration of recom-

binant Gqa from E. coli and Gqa expressed in Sf9 membranes

is explained by the fact that bacterially expressed G proteins

do not possess covalent modifications (Graziano et al. 1987,

1989) that have an impact on electrophoretic mobility. The

intensity of the immunoreactive band with 0.2 lg of

recombinant mammalian Gqa (equivalent to �5 pmoles of

protein) is �five-fold lower than the intensity of the Gqa

band observed with 50 lg of protein from Sf9 membranes

expressing hH2R plus Gqa. Thus, we estimated that in Sf9

membranes Gqa is expressed at levels of �500 pmol/mg.

These levels clearly surpass the expression levels achieved

for GsaL and Gia2 in Sf9 membranes (Seifert et al. 1998a;

Wenzel-Seifert et al. 1999; Seifert and Wenzel-Seifert 2001).

The expression level of hH2R-Gqa was below the detection

limit of Ig 371754 (Fig. 3d). Similar difficulties in detecting

the Gqa moiety were previously reported for the b2AR-Gqa

fusion protein (Wenzel-Seifert and Seifert 2000). Although

the analysis of the hH2R-Gqa fusion protein with the M1

monoclonal antibody suggested the expression of both

coupling partners in the fusion protein (Fig. 1c), we wished

to detect the Gqa moiety as well. Therefore, we reverse-

transcribed mRNA of Sf9 cells infected with the hH2R-Gqa

baculovirus and amplified the Gqa sequence by PCR. As was

reported for the b2AR-Gqa fusion protein (Wenzel-Seifert

and Seifert 2000), a 1146-bp sequence was amplified from

hH2R-Gqa-infected Sf9 cells that contained an Eco RI site at

position 937 and a Pst I site at position 1063 (data not

shown).

Taken together, the radioligand binding and immunoblot

studies demonstrate that hH2R, hH2R-GsaS, hH2R-Gqa, GsaS

and Gqa were all expressed in Sf9 cells, although at different

levels. By analogy to data obtained for Gi-proteins (Wenzel-

Seifert et al. 1999; Seifert and Wenzel-Seifert 2001), the

large molar excess of Gqa relative to hH2R was expected to

provide excellent conditions for efficient GPCR/G protein

coupling, particularly for ternary complex formation. Our

failure to detect the endogenous Gs- and Gq-proteins of Sf9

cells with various antibodies could be due to substantial

structural differences between mammalian and insect cell

G proteins and/or low expression levels of endogenous

G proteins. We cannot discriminate between these two

possibilities, but the efficient activation of AC by b2AR,

hH2R, gpH2R and rH2R and the RGS protein-enhancement

of PAFR-, hH1R- and gpH1R-stimulated GTP hydrolysis (see

below) provided functional evidence for the presence of

endogenous Gs- and Gq-proteins in the insect cells.

Ternary complex formation

One of the earliest steps of the G protein cycle is the

formation of the ternary complex (Kent et al. 1980). The

ternary complex consists of the agonist-occupied GPCR and

the guanine nucleotide-free G protein and is characterized by

high agonist-affinity. In most systems, binding of the

hydrolysis-resistant GTP analog GTPcS disrupts the ternary

complex, thereby decreasing agonist-affinity of GPCR.

However, in several systems comprising GPCRs coupled to

various classes of G proteins including Gi/Go-proteins that

exchange guanine nucleotides rapidly (Gilman 1987),

GTPcS does not or only partially disrupt the ternary

complex, indicative of tight GPCR/G–protein interaction

even in the presence of GTPcS (Stiles 1985; Childers et al.

1993; Szele and Pritchett 1993; Gurdal et al. 1997; Seifert

et al. 1998a). Similar to the b2AR (Seifert et al. 1998a,b),

ternary complex formation with hH2R was assessed

(a) (b)

(c) (d)

Fig. 3 Analysis of the expression of Gqa and hH2R-Gqa in Sf9

membranes. Sf9 membranes expressing various proteins were



the anti-Gqa Igs 371730 (a), 371752 (b) or 371754 (c and d). Shown

are the horseradish peroxidase-reacted membranes. Numbers on the

left of membranes designate masses of marker proteins in kDa. In

(a–c), 100 lg of membrane protein from Sf9 cells and 0.2 lg of

purified recombinant Gqa (rec. Gqa) were loaded onto each lane. In (d),

the amounts of protein loaded onto each lane are given below the

membrane.



indirectly by analyzing the displacement of antagonist

binding by unlabeled HIS.

With respect to b2AR/Gs interaction, it has been shown

that the coexpression of mammalian bc-complexes is not

required for ternary complex formation, presumably because

endogenous insect bc-complexes can interact with the b2AR

and mammalian Gsa (Seifert et al. 1998a). In addition, the

presence of mammalian bc-complexes renders the detection

of agonist-stimulation of AC more difficult (Seifert et al.

1998a). Therefore, we did not coexpress bc-complexes in

hH2R/Gsa reconstitutions. However, mammalian bc-com-

plexes enhance GPCR/G protein coupling in terms of ternary

complex formation (Seifert et al. 1998a), and mammalian

bc-complex is required to prevent aggregation of Gqa in Sf9

cells (Hepler et al. 1993). Based on these considerations we

coexpressed mammalian b1c2-complex in the hH2R/Gqa

reconstitutions in order to provide optimal coupling condi-

tions. As reported before (Wenzel-Seifert et al. 1998),

expression of b1c2-complex was associated with decreased

expression of insect cell bc-complex as assessed by immu-

noblotting with the anti-Gb-common Ig (data not shown).

Figure 4 shows the competition isotherms for HIS at the

various constructs in the absence and presence of GTPcS,

and Table 1 provides a summary of the non-linear regression

analysis of the competition isotherms. In membranes

expressing hH2R, HIS inhibited [3H]TIO binding with a

steep monophasic curve, and GTPcS did not shift the

HIS-competition curve to the right (Fig. 4a and Table 1).

These data indicate that ternary complex formation of hH2R

with endogenous Gs- and/or Gq-proteins is poor and/or below

the detection limit of the assay, despite the fact that the

expression level of hH2R was relatively low (�1 pmol/mg as

assessed by immunoblotting). In agreement with the data for

hH2R, the b2AR is also unable to form measurable ternary

complexes with insect cell G proteins, even when expressed

at low levels (Seifert et al. 1998a).

Although there was only a �1 : 1 ratio of hH2R and GsaS

coexpressed in Sf9 membranes (Fig. 1a and 2a), hH2R and

GsaS formed a ternary complex as is evident from the shallow

HIS-competition curve and the right-shift of the curve by

GTPcS (Fig. 4b and Table 1). Non-linear regression analysis

revealed that in the absence of GTPcS, �40% of the hH2Rs

formed a ternary complex with GsaS. These data clearly

indicate that a large fraction of the expressed hH2R and GsaS

molecules are localized within the same membrane micro-

compartment and efficiently interact with each other.

Fig. 4 Competition by HIS of [3H]TIO binding in Sf9 membranes

expressing hH2R without or with various G protein subunits and in

membranes expressing hH2R-Ga fusion proteins. [3H]TIO binding in

Sf9 membranes was performed as described in Materials and meth-

ods. Reaction mixtures contained membranes expressing the proteins

indicated in (a–e), 10 nM [3H]TIO and HIS at increasing concentra-

tions. Tubes additionally contained solvent (control) or GTPcS

(10 lM). Data points are the means ± SD of 3–7 independent exper-

iments performed in duplicates. Data were analyzed by non-linear

regression for best fit to one-site or two-site competition curves. The

expression levels of hH2R and GsaS were �1 pmol/mg each (see

Figs 1a and 2a), the expression level of hH2R-GsaS was �2 pmol/mg

(see Fig. 1b), the expression level of hH2R-Gqa was �1 pmol/mg (see

Fig. 1c), and the expression level of Gqa was �500 pmol/mg (Fig. 3d).



For the b2AR, it has been shown that ternary complex

formation in a b2AR-Gsa fusion protein is more efficient than

in a coexpression system of b2AR plus Gsa (Seifert et al.

1998a). Similarly, in the hH2R-Gsa fusion protein, a larger

fraction of hH2Rs (�60% vs. �40% in the coexpression

system) formed a ternary complex in the absence of GTPcS(Figs 4b and c and Table 1). In the membranes expressing

hH2R-GsaS, GTPcS shifted the HIS-competition curve to the

right, but the agonist-competition curve was still shallow, i.e.

in the presence of GTPcS, �20% of the hH2Rs formed

a ternary complex. The partial GTPcS-insensitivity of high-

affinity agonist binding points to a very tight interaction of

hH2R with GsaS. GTPcS–insensitive interactions of GPCRs

with G proteins have been repeatedly observed (Citri and

Schramm 1982; Stiles 1985; Matesic et al. 1989; Childers

et al. 1993; Szele and Pritchett 1993; Gurdal et al. 1997;

Seifert et al. 1998a; Kelley et al. 2001). Given the only

incomplete labeling of hH2R constructs, specifically H2R-Ga

fusion proteins with [3H]TIO (see above), and the partially

GTPcS-insensitive high-affinity binding of HIS at hH2R-

GsaS it would have been desirable to study high-affinity

binding directly with [3H]HIS. Unfortunately, the Kh values

for HIS in membranes expressing hH2R + GsaS and hH2R-

GsaS are even higher than the Kd values for [3H]TIO

(Table 1), limiting assay sensitivity, and the interpretation of

[3H]HIS binding studies to H2R is complex, too (Steinberg

et al. 1985a,b,c).

Despite the favorable GPCR/G protein ratio in membranes

expressing hH2R plus Gqa compared to membranes exp-

ressing hH2R plus GsaS, the HIS-competition curves in the

hH2R/Gq coexpression system and hH2R-Gqa fusion protein

were steep and monophasic, and GTPcS shifted the compe-

tition curves only minimally to the right (Figs 4d and e).

These data show that Gqab1c2 is inefficient at stabilizing a

ternary complex with hH2R. It might be argued that

our inability to detect ternary complex formation in the

hH2R/Gqa systems is due to the fact that [3H]TIO labels only

a fraction of the expressed hH2R molecules and that the

molecules not labeled with [3H]TIO actually form a ternary

complex. However, with the non-fused hH2R, the [3H]TIO

saturation binding assay underestimates the actual expression

level to a much lesser extent than with hH2R-Gqa, and the

studies assessing guanine nucleotide exchange ([35S]GTPcSbinding and GTP hydrolysis) were consistent with the

agonist-competition studies (see below).

The H1R is a classic Gq-coupled GPCR (Leurs et al. 1995;

Hill et al. 1997). A typical property of Gq-coupled GPCRs

is the very long third intracellular loop (Wess 1997), a

feature that is also present in the H1R (Leurs et al. 1995;

Hill et al. 1997). Therefore, we expected to observe highly

efficient ternary complex formation in membranes expressing

hH1R or gpH1R expressed at 4.0 ± 0.4 pmol/mg and

3.5 ± 0.6 pmol/mg, respectively, as assessed by [3H]MEP

saturation binding plus �500 pmol/mg mammalian Gqa plus

b1c2-complex. However, similar to the data obtained with

membranes expressing hH2R plus Gqab1c2, the HIS compe-

tition isotherms for antagonist radioligand binding were steep

and monophasic in membranes expressing hH1R or gpH1R,

and GTPcS shifted the HIS competition curves only

minimally, if at all, to the right (data not shown). These

data indicated that Gqa, although expressed at high levels

in Sf9 membranes as assessed by immunoblotting and

expressed together with b1c2-complex to prevent aggregation

(Hepler et al. 1993), is nonetheless largely inactive and

therefore compromises the proper interpretation of data.

[35S]GTPcS binding studies

The [35S]GTPcS binding assay is a widely employed method

for analyzing GPCR-mediated guanine nucleotide exchange

at G proteins. In membranes expressing hH2R alone, HIS

had no stimulatory effect on [35S]GTPcS binding (Fig. 5a).

The inclusion of GDP into reaction mixtures may facilitate

the detection of agonist-stimulated [35S]GTPcS binding by

loading G proteins with GDP and enhancing the ability of

GPCR at promoting GDP dissociation (Wieland and Jakobs

1994). As predicted, GDP competed with [35S]GTPcS and

reduced basal [35S]GTPcS binding but did not unmask a

stimulatory effect of HIS on [35S]GTPcS binding in mem-

branes expressing hH2R alone (Fig. 5a). These data indicate

that hH2R couples to endogenous Gs- and/or Gq-proteins

only inefficiently and/or that the sensitivity of the binding

assay is too low to monitor coupling. In membranes

expressing hH2R plus GsaS, a small stimulation of

[35S]GTPcS binding by HIS was detected in the absence

and presence of GDP (Fig. 5b). In membranes expressing

hH2R-GsaS, HIS had a substantial stimulatory effect on

[35S]GTPcS binding in the absence and presence of GDP

(Fig. 5c). These data show that the agonist-occupied hH2R

efficiently enhanced GDP dissociation from, and [35S]-

GTPcS binding to, its fused GsaS partner. Similar data were

obtained for b2AR-Gsa fusion proteins (Seifert et al. 1998a;

Wenzel-Seifert and Seifert 2000; Liu et al. 2001b).

In membranes expressing hH2R plus Gqa, agonist-stimu-

lation of [35S]GTPcS binding was minimal, regardless of the

absence or presence of GDP (Fig. 5d). In membranes

expressing hH2R-Gqa, HIS stimulated [35S]GTPcS binding

in the absence of GDP by 16%, but GDP did not further

increase the stimulatory effect of HIS on [35S]GTPcSbinding (Fig. 5e). We also conducted time course studies

of [35S]GTPcS binding to hH2R-Gqa using incubation times

between 5 and 180 min However, these studies failed to

reveal a more suitable time point than the 60 min incubation

used for the GDP competition experiments to reveal a larger

stimulatory effect of HIS on [35S]GTPcS binding (data not

shown).

Similarly to the situation concerning ternary complex

formation, we were unable to establish a system in which

efficient coupling of a GPCR to Gqa could be demonstrated



in terms of [35S]GTPcS binding. Specifically, in membranes

expressing hH1R, gpH1R or PAFR, the latter being a dually

Gi/Gq-coupled GPCR (Shimizu et al. 1992; Seifert 2001),

plus Gqab1c2, agonist-stimulations of [35S]GTPcS binding

were not greater than in membranes expressing hH2R plus

Gqab1c2 or hH2R-Gqa plus b1c2-complex (data not shown).

Similar results as for [35S]GTPcS binding were obtained

steady-state GTPase activity in membranes expressing hH1R,

gpH1R or PAFR plus Gqab1c2 (data not shown). These

findings support the results of the agonist binding studies

suggesting that the majority of the expressed Gqa molecules

is functionally inactive.

AC studies

The assessment of AC activity is the most sensitive

parameter for studying GPCR/Gs-protein coupling because

of the signal amplification at the effector level (Gilman 1987;

Birnbaumer et al. 1990). Indeed, although we obtained no

evidence for ternary complex formation of hH2R with, and

agonist-stimulation of [35S]GTPcS binding to, endogenous

Gs-proteins (Figs 4a and 5a), AC studies revealed that the

hH2R couples to insect Gs-proteins quite efficiently. In the

presence of GTP, HIS stimulated AC activity in Sf9

membranes expressing hH2R by almost five-fold, and this

AC activation even surpassed AC activation by GTPcS(Fig. 6a). Previous studies already showed that the b2AR and

Fig. 6 Regulation of AC activity in Sf9 membranes expressing hH2R,

hH2R plus GsaS and hH2R-GsaS fusion protein. AC activity in Sf9

membranes was determined as described in Materials and methods.

Reaction mixtures contained membranes expressing the proteins

indicated in (a–c) and solvent (basal), GTP (100 lM), GTP (100 lM)

plus HIS (100 lM) or GTPcS (10 lM). Data points are the means ± SD

of 3–6 independent experiments performed in triplicates. The exp-

ression levels of hH2R and GsaS were �1 pmol/mg each (see Figs 1a

and 2a), and the expression level of hH2R-GsaS was �2 pmol/mg (see

Fig. 1b).

Fig. 5 Effect of HIS on [35S]GTPcS binding in Sf9 membranes

expressing hH2R without or with various G protein subunits and in

membranes expressing hH2R-Ga fusion proteins. [35S]GTPcS binding

in Sf9 membranes was performed as described in Materials and

methods. Reaction mixtures contained membranes expressing the

proteins indicated in (a–e), 0.4 nM [35S]GTPcS and GDP at increasing

concentrations. Tubes additionally contained solvent (basal) or HIS

(100 lM). Data points are the means ± SD of 3–5 independent

experiments performed in triplicates. Data were analyzed by non-

linear regression and were best fitted to one-site competition curves.

The expression levels of hH2R and GsaS were �1 pmol/mg each (see

Figs 1a and 2a), the expression level of hH2R-GsaS was �2 pmol/mg

(see Fig. 1b), the expression level of hH2R-Gqa was �1 pmol/mg (see

Fig. 1c), and the expression level of Gqa was �500 pmol/mg (Fig. 3d).



rH2R activate AC via Sf9 cell Gs-proteins (Parker et al.

1991; Chidiac et al. 1994; Kuhn et al. 1996; Beukers et al.

1997). In Sf9 membranes expressing gpH2R at 0.25 pmol/mg

as assessed by [3H]TIO saturation binding, HIS increased

AC activity from 0.95 ± 0.10 pmol/mg/min to 1.90 ± 0.15

pmol/mg/min (mean ± SD, n ¼ 3). In Sf9 membranes

expressing rH2R at 0.18 pmol/mg as assessed by

[3H]TIO saturation binding, HIS increased AC activity

from 1.03 ± 0.08 pmol/mg/min to 1.95 ± 0.22 pmol/mg/min

(mean ± SD, n ¼ 3). Collectively, these data show that the

AC assay possesses sufficiently high sensitivity to demon-

strate coupling of various mammalian GPCRs and of H2R

species isoforms to insect cell Gs-proteins.

We also wished to determine how efficient the coupling

of mammalian GPCRs to insect cell Gs-proteins is relative

to the coupling of insect cell GPCRs to Gs-proteins. To

address this question, we examined the effect of octopamine

(10 nM – 1 mM) on AC activity in membranes from

uninfected Sf9 cells. Octopamine is an important neuro-

transmitter in insect cells (Evans and Robb 1993) and was

reported to efficiently activate AC in membranes from

uninfected Sf9 cells (Orr et al. 1992). However, although

we could readily reproduce the reported AC activation by

forskolin (data not shown) and activate AC in membranes

from uninfected Sf9 cells with GTPcS (Seifert et al.

1998a), we were unable to demonstrate stimulatory effects

of octopamine on AC activity in this system (data not

shown). We do not have a satisfying explanation for these

apparent discrepancies, but differences in the cell culture

conditions may be a factor (see Materials and methods)

(Orr et al. 1992). Thus, unfortunately, we cannot assess

how efficient coupling of mammalian GPCRs to insect

Gs-proteins is relative to coupling of insect cell GPCRs to

those Gs-proteins.

In Sf9 membranes expressing hH2R plus GsaS, GTP

per se had a substantial stimulatory effect on AC activity

(Fig. 6b), presumably reflecting the constitutive, i.e. agonist-

independent, activity of the hH2R (Smit et al. 1996). HIS

increased AC activity in membranes coexpressing hH2R

and GsaS by �55%. Compared to membranes expressing

hH2R alone, the absolute AC activities in membranes

expressing hH2R plus GsaS were considerably higher, but in

terms of relative agonist-stimulation, coupling of hH2R to

insect cell Gs-proteins actually surpassed hH2R-coupling to

mammalian GsaS. Compared with membranes coexpressing

hH2R and GsaS, the absolute AC activities, particularly

basal AC activity in the absence of GTP, in membranes

expressing hH2R-GsaS were all higher (compare Figs 6b

and c). These data corroborate the data obtained for b2AR-Gsa fusion proteins showing that fusion of a GPCR to Gsa

increases the absolute AC activities (Bertin et al. 1994;

Seifert et al. 1998a). Presumably, fusion proteins and AC

molecules are targeted to the same microcompartments,

forming highly efficient AC-activating transducisomes

(Seifert et al. 1999).

Ca2+ signaling studies

For the analysis of Gq-signaling, we took advantage of the

fact that in many systems including Sf9 cells, PLC activation

results in Ca2+ mobilization from intracellular stores with

subsequent Ca2+ influx from the extracellular space through

cation channels (Hu et al. 1994; Harteneck et al. 1995; Kuhn

et al. 1996). Initially we wished to answer the question

whether there is an endogenous GPCR-regulated Ca2+

signaling pathway in Sf9 cells. We focused our attention

on octopamine and nucleotides since insects are known to

possess GPCRs for these signaling molecules (Galun et al.

1988; Evans and Robb 1993). Rises in [Ca2+]i were

monitored with the Ca2+-sensitive dye fura-2. However,

octopamine, ATP, GTP, ITP, XTP, UTP, CTP, ADP and UDP

(10 nM – 1 mM) did not exhibit stimulatory effects on [Ca2+]iin uninfected Sf9 cells. Thus, similar to the situation with

GPCR/Gs-protein coupling, we could not identify intact

GPCR/Gq-protein coupling with endogenous GPCRs in Sf9

cells.

Given this situation, we then explored whether Ca2+

signaling can be activated by stimuli that circumvent GPCRs.

We used TG, a Ca2+ ATPase inhibitor that activates Ca2+

influx from the extracellular space via depletion of intracel-

lular Ca2+ stores (Thastrup et al. 1994), and the wasp venom

MP. MP and related peptides exhibit pleiotropic effects such

as activation of Gi/Go-proteins (Higashijima et al. 1988),

nucleoside diphosphokinase (Klinker et al. 1994) and Ca2+

influx mechanisms (Suh et al. 1996; Hirata et al. 2000). In a

concentration-dependent manner, TG and MP induced large

and sustained increases in [Ca2+]i in uninfected Sf9 cells

(Figs 7a and b). These rises in [Ca2+]i were largely due to

influx from the extracellular space since in the absence of

extracellular Ca2+, the rises in [Ca2+]i were strongly dimin-

ished (Table 2). The lanthanides Gd3+ and La3+ (10 lM each)

had been shown to inhibit the endogenous cation entry

pathway in Sf9 cells (Hu et al. 1994). In agreement with

these findings, Gd3+ and La3+ (10 lM each) inhibited TG- and

MP-induced rises in [Ca2+]i in Sf9 cells (Table 2). Thus,

Ca2+ signaling in Sf9 cells can be activated at least at a

postreceptor level.

Next, we set out to study the effects of mammalian GPCRs

in Sf9 cells. Previous studies had shown that detection of

agonist-stimulated rises in [Ca2+]i in Sf9 cells is dependent

on the length of the infection period. Specifically, GPCR-

mediated rises in [Ca2+]i were only observed in the early

stages of infection (24–36 h after addition of virus) but not in

the later stages of infection (Kuhn et al. 1996; Kukkonen

et al. 1996; Leopoldt et al. 1997). In Sf9 cells that had been

infected for 24 h with baculovirus encoding hH2R at a

1 : 100 dilution, TG (10 lM) induced a long-lasting increase



in [Ca2+]i as it did in uninfected cells (compare Figs 7a and

c), but the magnitude of the rise in [Ca2+]i decreased from

354 ± 58 nM to 101 ± 18 nM (mean ± SD, n ¼ 15). These

data indicate that the expression of some distal components

of the Ca2+ signaling pathway, such as transmembrane cation

channels, are strongly reduced as the result of baculovirus

infection. We failed to detect any HIS-induced rises in

[Ca2+]i in Sf9 cells using hH2R alone, hH2R plus Gqa without

and with b1c2 and hH2R-Gqa without and with b1c2 at virustiters of 1 : 100–1: 10 000 and using infection periods

ranging from 16 h to 48 h (Fig. 7c and data not shown).

Moreover, extensive studies with rH2R, gpH2R, PAFR, hH1R

and gpH1R using different virus dilutions and infection times

failed to unmask any stimulatory effect of GPCR agonists on

[Ca2+]i in Sf9 cells, regardless of whether GPCRs were

studied alone or with mammalian Gqab1c2 (data not shown).Our inability to detect a stimulatory effect of GPCRs on

[Ca2+]i in Sf9 cells cannot be explained by an increase in

basal [Ca2+]i as a result of baculovirus infection, i.e. basal

[Ca2+]i in infected Sf9 cells ranged between �60–120 nM

(see legends to Fig. 7 and Table 2 and data not shown). We

can also rule out the possibility that baculovirus infection

resulted in a general defect of GPCR/Gq-protein coupling

since the below-described GTPase studies with RGS proteins

demonstrated intact GPCR-regulation of guanine nucleotide

exchange at insect cell Gq-proteins even after an infection

period of 48 h. Most likely, the reduced expression of distal

components of the signaling pathway that are of importance

for activation of Ca2+ entry by TG and GPCRs account for

our failure to detect GPCR-mediated rises in [Ca2+]i in Sf9

cells. Collectively, in our hands, the fura-2 assay proved to be

unsuitable to study GPCR/Gq-protein coupling, be it endog-

enous GPCRs and G proteins or mammalian GPCRs and

G proteins.

Our disappointing results regarding GPCR-regulation of

rises in [Ca2+]i in Sf9 cells contrast to reports showing

GPCR-mediated rises in [Ca2+]i in those cells (Harteneck

et al. 1995; Kuhn et al. 1996; Kukkonen et al. 1996;

Leopoldt et al. 1997). We can exclude insensitivity of

the fura-2 method for the discrepancies since basal

[Ca2+]i-values in uninfected and infected Sf9 cells were

low (legends to Fig. 7 and Table 2 and data not shown), dye

loading was efficient and dye leakage was low (data not

shown). Moreover, we can exclude the possibility that the

lack of stimulatory effect of HIS with the hH2R is a

peculiarity of this GPCR species isoform since the gpH2R

and rH2R were ineffective in the fura-2 assay as well, but

they were active in the AC assay. The same rH2R baculovirus

that was effective at reconstituting Gq- and Gs-coupling in

Fig. 7 Effects of TG, MP and HIS on [Ca2+]i in Sf9 cells. [Ca2+]i in Sf9

cells was determined as described in Materials and methods. Super-

imposed original fura-2 fluorescence recordings are shown. (a) and (b)

show recordings from uninfected Sf9 cells, and (c) shows recordings

from Sf9 cells that had been infected with hH2R baculovirus (1 : 100)

for 24 h. The arrows indicate the addition of stimuli at the concentra-

tions shown at the individual recordings. Basal [Ca2+]i in uninfected

Sf9 cells was 95 ± 13 nM (mean ± SD, n ¼ 16). Basal [Ca2+]i in hH2R

baculovirus-infected cells was 87 ± 18 nM (mean ± SD, n ¼ 6). The

Figure shows relative fluorescence changes. Absolute increases in

[Ca2+]i induced by TG and MP in uninfected and infected Sf9 cells are

reported in Table 2 and in the text.

Table 2 Effects of Gd3+, La3+ and EGTA on TG- and MP-induced

rises in [Ca2+]i in uninfected Sf9 cells

Addition

Increase in [Ca2+]i (nM)

TG (10 lM) MP (30 lM)

Solvent (control) 354 ± 58 198 ± 5

GdCl3 (10 lM) 92 ± 54 105 ± 24

LaCl3 (10 lM) 93 ± 16 104 ± 17

EGTA (1 mM) 24 ± 10 45 ± 10

Rises in [Ca2+]i in uninfected Sf9 cells were determined as described

in Materials and methods. Cuvettes contained either solvent (control)

or lanthanides in the presence of 1 mM CaCl2 or 1 mM EGTA in the

absence of CaCl2. After monitoring basal [Ca2+]i for 1 min, TG or MP

at the concentrations shown in the table were added to cuvettes. The

maximum stimulus-induced rises in [Ca2+]i after 3 min were calcu-

lated. Basal [Ca2+]i in uninfected Sf9 cells was 93 ± 13 nM. Data

shown are the means ± SD from three to seven experiments.



previous studies (Harteneck et al. 1995; Kuhn et al. 1996;

Leopoldt et al. 1997), in our hands reconstituted only

Gs-coupling. In addition, gpH1R expressed in Sf9 cells

reconstituted rises in [Ca2+]i in previous studies (Harteneck

et al. 1995; Kuhn et al. 1996; Leopoldt et al. 1997) but not

in our hands. Thus, it is evident that coupling of different

GPCRs to insect Gs-proteins in Sf9 cells is very reproducible

among various laboratories, whereas Gq-coupling is not

readily reproduced. Presently, we do not have a satisfactory

explanation for the different results from various laboratories

but differences in cell culture conditions could have an as yet

unappreciated impact on GPCR/Gq-protein coupling (see

Materials and methods) (Harteneck et al. 1995; Kuhn et al.

1996; Leopoldt et al. 1997).

GTPase studies

In view of the unsatisfying results regarding the expression

of functionally active mammalian Gqa and the difficulties to

study Ca2+ signaling in Sf9 cells we aimed at establishing

another sensitive method to study at least coupling of

mammalian GPCRs to insect cell Gq-proteins. We considered

five findings. First, it has been shown that RGS proteins can

dramatically enhance the agonist-stimulated steady-state

GTP hydrolysis of Gq-coupled GPCRs (Ingi et al. 1998;

Mukhopadhyay and Ross 1999). Second, the RGS proteins

identified so far do not act on Gs-proteins (Ross and Wilkie

2000). Third, although the RGS proteins studied herein

(RGS4 and GAIP) are not specific for Gq-proteins but also

act on Gi-proteins (Ross and Wilkie 2000), this does not

constitute a problem for our purposes since is has been very

well documented by various laboratories including our own

that Sf9 cells do not express Gi-proteins (Quehenberger et al.

1992; Obosi et al. 1996; Leopoldt et al. 1997; Wenzel-

Seifert et al. 1998; Kuhn and Gudermann 1999). Fourth, the

analysis of HIS-stimulated [35S]GTPcS binding and steady-

state GTP hydrolysis in Sf9 membranes expressing hH1R or

gpH1R plus Gia2b1c2 confirmed that coupling of those classic

Gq-coupled GPCRs to Gi-proteins is very poor (data not

shown). Fifth, the basal steady-state GTPase activity in Sf9

cell membranes is low, providing an excellent signal-to noise

ratio (Seifert et al. 1998a; Wenzel-Seifert et al. 1999). Based

on these considerations, we reasoned that the analysis of the

effects of RGS proteins on GTP hydrolysis catalyzed by

GPCRs coupled to insect cell G proteins could provide a

sensitive method to study GPCR/Gq-protein coupling pro-

vided that the mammalian RGS proteins would interact with

the insect cell Gq-proteins.

Before conducting the GTPase experiments, we had to

consider two other important factors. First, it has been shown

that various Gq-coupled GPCRs interact differentially with

RGS proteins (Xu et al. 1999). To address this point, we

combined RGS proteins from different families (RGS4, RGS

protein family R4; GAIP, RGS protein family RZ) (Ross and

Wilkie 2000) with GPCRs from different families (H2R and

H1R, GPCR family 1a; PAFR, GPCR family 1b) (Bockaert

and Pin 1999). It should be noted that only the H1R, but not

the H2R and PAFR possess the large third intracellular loop

that is typical for Gq-coupled GPCRs (Shimizu et al. 1992;

Leurs et al. 1995; Hill et al. 1997; Wess 1997). Second, it is

emerging that there are significant functional differences

between species isoforms of HxR subtypes including the H2R

(Ligneau et al. 2000; Lovenberg et al. 2000; Kelley et al.

2001; Liu et al. 2001a). Therefore, we compared the human

and guinea pig isoforms both of the H1R and H2R.

Figure 8a shows an immunoblot with the anti-FLAG Ig of

Sf9 membranes expressing the b2AR, PAFR and hH2R

without or with RGS4. The intensities of the GPCR bands

were similar regardless of whether RGS4 was expressed or

not. Similar data concerning expression were obtained upon

(a)

(b)

Fig. 8 Analysis of the expression of hH2R, b2AR, PAFR and RGS4 in

Sf9 membranes. Sf9 membranes expressing various proteins were



the anti-FLAG Ig (M1 antibody) (a) or anti-RGS4 Ig (b). Shown are the

horseradish peroxidase-reacted membranes. Numbers on the left

designate masses of marker proteins. On each lane, 50 lg of protein

were applied. The b2AR was expressed at a level of 3.5 pmol/mg.

– and + in (a) and (b) designate the absence or presence, respectively,

of RGS4 baculovirus during the Sf9 cell culture. hH2R and hH2R*

designate two different sets of membrane preparations expressing

hH2R. unif., uninfected Sf9 cells.



coexpression of GPCRs with GAIP (data not shown). These

data show that expression of RGS proteins does not interfere

with the expression of GPCRs, providing precisely defined

experimental conditions for studying the effect of RGS

proteins on GPCR/G protein coupling.

RGS4 expressed in HEK293 cells possesses a molecular

mass of �32 kDa (Huang et al. 1997). In agreement with

those data, RGS4 expressed in Sf9 cell membranes migrated

as a �32 kDa protein (Fig. 8b). In membranes from

uninfected Sf9 cells or Sf9 cells infected with GPCR

baculovirus only, the 32-kDa band was absent. In all

membrane preparations, an additional �40-kDa band was

observed. This band presumably represents a non-specific

reaction of the anti-RGS4 Ig. Upon infection with baculo-

viruses, the 40-kDa band became less intense, indicating

decreased expression of this protein in infected Sf9 cells.

For the hH1R and gpH1R, we determined expression levels

with the [3H]MEP saturation binding assay. Again, RGS

proteins did not largely change H1R expression levels. The

expression levels of hH1R ± RGS4 or GAIP ranged between

4.0 and 4.9 pmol/mg, and the expression levels of

gpH1R ± RGS proteins ranged between 3.5 and 4.5 pmol/mg.

Figure 9a shows that RGS4 could also be coexpressed with

hH1R and gpH1R. Similar to RGS4, GAIP migrates as a

�32-kDa protein when expressed in Sf9 membranes

(Fig. 9b). The molecular mass of GAIP in Sf9 membranes

corresponds with the data obtained for GAIP expressed in

HEK293 cells (Huang et al. 1997). There was no immuno-

reactivity with the anti-GAIP Ig in membranes from

uninfected Sf9 cells. Collectively, these data show that

uninfected Sf9 cells do not express mammalian-type RGS4

and GAIP, that RGS4 and GAIP can be efficiently expressed

in Sf9 membranes and that RGS protein expression has little

effect on GPCR expression levels.

In membranes expressing the hH2R, b2AR, PAFR, gpH2R,

hH1R and gpH1R alone, the agonist-stimulations of steady-

state GTP hydrolysis were small (Fig. 10a–c). RGS4

enhanced basal and agonist-stimulated GTP hydrolysis in

membranes expressing PAFR, hH1R and gpH1R (Figs 10a

and b). In the presence of RGS4, the stimulatory effects of

agonist in the membranes expressing PAFR amounted to

78%, and the stimulatory effects of agonist in membranes

expressing hH1R and gpH1R amounted to 142% and 174%,

respectively. These data suggest that RGS4 enhances H1R

agonist-stimulated GTP hydrolysis more efficiently than

(a) (b)

Fig. 9 Analysis of the expression of RGS4 and GAIP in Sf9 mem-

branes. Sf9 membranes expressing various proteins were prepared,

separated by SDS–PAGE on gels containing 10% (w/v) acrylamide,

transferred onto Immobilon P membranes and probed with the anti-

RGS4 Ig (a) or anti-GAIP Ig (b). Shown are the horseradish peroxi-

dase-reacted membranes. Numbers on the left designate masses of

marker proteins. On each lane, 50 lg of protein were applied. Cell

cultures contained either RGS4 baculovirus, GAIP baculovirus or no

virus (uninfected). Cell cultures containing RGS protein baculoviruses

also contained the HxR viruses provided below the gel.

Fig. 10 Effect of RGS4 and GAIP on basal and agonist-stimulated

GTPase activities in Sf9 membranes. GTPase activity in membranes

expressing various GPCRs without or with RGS4 (a and b) and without

or with GAIP (c) was determined as described in Materials and

methods. GTPase activity was determined under basal conditions b or

in the presence of agonist a. For membranes expressing hH1R, hH2R,

gpH1R and gpH2R ± RGS proteins, the agonist was HIS (100 lM). For

membranes expressing b2AR, the agonist was (–)-isoproterenol

(10 lM). For membranes expressing PAFR, the agonist was PAF

(10 lM). Data shown are the means ± SD of three independent ex-

periments performed in triplicates.



PAFR agonist-stimulated GTP hydrolysis. These data are in

accordance with the data of Xu et al. (1999), demonstrating

GPCR-specificity of the effects of RGS proteins. Our data are

also in agreement with previous findings showing that RGS4

can regulate PAFR signaling (Richardson et al. 2001) and

that RGS4 can accelerate GTP hydrolysis stimulated by

Gq-coupled GPCRs (Mukhopadhyay and Ross 1999).

In contrast to the data obtained with the PAFR and

H1Rs, RGS4 did not enhance the agonist-stimulated GTP

hydrolysis in membranes expressing the b2AR, hH2R and

gpH2R (Figs 10a and b). These data are compatible with the

notion that the b2AR and H2R are preferentially Gs-coupled

GPCRs and that RGS proteins do not enhance the GTPase

activity of Gs-proteins (Ross and Wilkie 2000). An expla-

nation of the higher basal GTPase activity in membranes

expressing hH2R than in membranes expressing b2AR could

be that the b2AR virus is more efficient than the hH2R virus at

suppressing the expression of endogenous G proteins. Thus,

the chances of detecting an enhancement of H2R-stimulated

GTP hydrolysis by RGS proteins should have been even

higher than for the b2AR.In view of the differential effects of RGS4 on agonist-

stimulated GTP hydrolysis in membranes expressing PAFR,

hH1R and gpH1R (Figs 10a and b), we also examined the

effects of GAIP, belonging to a different RGS protein family

than RGS4 (Ross and Wilkie 2000), on GTPase activity in

Sf9 membranes. Similar to the data obtained with RGS4,

GAIP strongly enhanced basal and agonist-stimulated GTP

hydrolysis in membranes expressing hH1R and gpH1R

(Fig. 10c). In the presence of GAIP, the agonist-stimulations

of GTP hydrolysis in membranes expressing hH1R and

gpH1R amounted to 126% and 140%, respectively. In

contrast, GAIP did not enhance basal and agonist-stimulated

GTP hydrolysis in membranes expressing hH2R (Fig. 10c).

Collectively, the comparison of GTPase activity without and

with RGS proteins was the only method with which we could

provide positive evidence for GPCR-coupling to endogenous

Gq-proteins. Using this approach, we did not detect efficient

coupling of the H2R to Gq-proteins.

Conclusions

In agreement with literature data on native and recombinant

systems (Bristow et al. 1982; Gespach and Abita 1982;

Mitsuhashi et al. 1989; Leurs et al. 1994; Alewijnse et al.

1998; Wang et al. 2000), our present data show that hH2R

robustly couples to insect cell and mammalian Gs-proteins.

By analogy to the b2AR (Seifert et al. 1998a), hH2R couples

to mammalian Gs-proteins more efficiently than to insect cell

Gs-proteins in terms of ternary complex formation and

[35S]GTPcS binding, and coupling of hH2R to mammalian

Gsa is more efficient in the fused than in the non-fused state.

Consistent with the literature, too, are our findings that the

H2R from other species, specifically rat and guinea pig,

couples to Gs-proteins as well (Traiffort et al. 1992b; Leurs

et al. 1995; Kuhn et al. 1996; Beukers et al. 1997; Hill et al.

1997; Leopoldt et al. 1997).

In contrast to the data on Gs-proteins, the data on

Gq-coupling of hH2R are less straightforward to interpret.

Specifically, we were unable to reconstitute efficient coupling

of classic Gq-coupled GPCRs such as hH1R and gpH1R with

mammalian Gqa in terms of ternary complex formation,

[35S]GTPcS binding and GTP hydrolysis, although Gqa is

highly expressed as assessed by immunoblotting. Thus, we

cannot rule out the possibility that the poor coupling of hH2R

to fused and coexpressed mammalian Gqa in Sf9 cells is

attributable to functional inactivity of the expressed Gqa. As

another complication, we were unable to demonstrate

GPCR/Gq-protein coupling using the fura-2 assay that is

a sensitive and widely used method to analyze

GPCR/Gq-protein coupling. In our hands, the only useful

and sensitive method to study coupling of GPCRs to

Gq-proteins was the steady-state GTPase assay that takes

advantage of the fact that Sf9 cells are devoid of Gi-proteins

and that RGS-proteins enhance agonist-stimulated steady-

state GTP hydrolysis catalyzed by Gq-coupled GPCRs.

Using this methodology, we showed that RGS proteins from

two different families enhance agonist-stimulated GTP

hydrolysis catalyzed by various Gq-coupled GPCRs includ-

ing hH1R, whereas RGS proteins did not enhance GTP

hydrolysis stimulated by the hH2R and another classic

Gs-coupled GPCR, the b2AR. Thus, despite significant

limitations regarding the analysis of mammalian Gq-proteins

and assessment of Gq-mediated activation of effector

systems, we can nonetheless make the statement that the

hH2R couples more efficiently to insect cells Gs-proteins than

to insect cell Gq-proteins.

On first glance, our present data seem to be in contrast

to previous reports demonstrating coupling of hH2R to

Gq-proteins as assessed by rises in [Ca2+]i (Mitsuhashi et al.

1989; Seifert et al. 1992; Burde and Seifert 1996). However,

in agreement with our present results, hH2R fails to couple to

Gq-proteins when expressed in Chinese hamster ovary cells

(Leurs et al. 1994). Intriguingly, even in systems in which

Gq-coupling of hH2R was observed, the stimulatory effects

of hH2R on [Ca2+]i were small compared to a prototypical

Gq-coupled GPCR, namely the P2U-receptor (Seifert et al.

1992). Thus, depending on the specific system studied,

inefficient hH2R-coupling to Gq-proteins may or may not

reach the threshold to result in rises in [Ca2+]i, whereas the

robust hH2R-coupling to Gs always reaches the threshold to

result in AC activation. Similar to hH2R, rH2R has been

consistently reported to mediate AC activation, whereas the

data concerning coupling of rH2R to Gq-proteins are

controversial (Traiffort et al. 1992a; Harteneck et al. 1995;

Kuhn et al. 1996; Beukers et al. 1997; Leopoldt et al. 1997).



Collectively, considering the literature data and our present

results it is likely that coupling of the hH2R and of the H2R

from other species to Gq-proteins is of little physiological

importance. Future studies aiming at elucidating the role of

the H2R in the regulation of brain functions should therefore

focus on Gs-mediated signaling.

Acknowledgements

The authors would like to thank M. T. Kelley for constructing the gpH2R

baculovirus and conducting some [3H]TIO saturation binding studies and

AC studies on gpH2R and the Reviewers of this paper for their most

valuable critique and suggestions. This work was supported by a New

Faculty Award of The University of Kansas, the National Institutes of

Health COBRE award 1 P20 RR15563 and matching support from the

State of Kansas and the University of Kansas and the National Institutes

of Health-Center for Research Resources grant ÔSchool of Pharmacy

Student Research ProgramÕ R25 RR10150.

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