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The Journal of Neuroscience, June 1, 1998, 18(11):4119-4132
Modulation of Rat Rotational Behavior by Direct Gene Transfer of
Constitutively Active Protein Kinase C into Nigrostriatal Neurons
Song
Song1,
Yaming
Wang1,
Sun-Yung
Bak1,
Matthew J.
During2,
John
Bryan1,
Oliver
Ashe1,
Donna B.
Ullrey1,
Laura E.
Trask1,
Frederick D.
Grant1,
Karen L.
O'Malley3,
Heimo
Riedel4,
David S.
Goldstein5,
Kim A.
Neve6,
Gerald J.
LaHoste7,
John F.
Marshall7,
John W.
Haycock8,
Rachael L.
Neve9, 10, and
Alfred I.
Geller1, 10
1 Division of Endocrinology, Children's Hospital,
Boston, Massachusetts 02115, 2 Departments of Surgery and
Medicine, Yale University School of Medicine, New Haven, Connecticut
06510, 3 Department of Anatomy and Neurobiology, Washington
University School of Medicine, St. Louis, Missouri 63110, 4 Department of Biological Sciences, Wayne State
University, Detroit, Michigan 48202, 5 Clinical
Neuroscience Branch, National Institute of Neurological Disorders and
Stroke, National Institutes of Health, Bethesda, Maryland 20892, 6 Research Service, Veterans Affairs Medical Center,
Portland, Oregon 97201, 7 Department of Psychobiology,
University of California, Irvine, California 92689, 8 Department of Biochemistry, Louisiana State University
Medical Center, New Orleans, Louisiana, 70119, 9 Molecular
Neurogenetics Laboratory, McLean Hospital, Belmont, Massachusetts
02178, and 10 Program in Neuroscience, Harvard Medical
School, Boston, Massachusetts 02115
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ABSTRACT |
The modulation of motor behavior by protein kinase C (PKC)
signaling pathways in nigrostriatal neurons was examined by using a
genetic intervention approach. Herpes simplex virus type 1 (HSV-1) vectors that encode a catalytic domain of rat PKC II (Pkc ) were developed. Pkc exhibited a constitutively active protein kinase activity with a substrate specificity similar to that of rat brain PKC.
As demonstrated in cultured sympathetic neurons, Pkc caused a
long-lasting, activation-dependent increase in neurotransmitter release. In the rat brain, microinjection of HSV-1 vectors that contain
the tyrosine hydroxylase promoter targeted expression to dopaminergic
nigrostriatal neurons. Expression of pkc in a small percentage of
nigrostriatal neurons (~0.1-2%) was sufficient to produce a
long-term ( 1 month) change in apomorphine-induced rotational
behavior. Nigrostriatal neurons were the only catecholaminergic neurons
that contained Pkc, and the amount of rotational behavior was
correlated with the number of affected nigrostriatal neurons. The
change in apomorphine-induced rotational behavior was blocked by a
dopamine receptor antagonist (fluphenazine). D2-like
dopamine receptor density was increased in those regions of the
striatum innervated by the affected nigrostriatal neurons. Therefore,
this strategy enabled the demonstration that a PKC pathway or PKC
pathways in nigrostriatal neurons modulate apomorphine-induced
rotational behavior, and altered dopaminergic transmission from
nigrostriatal neurons appears to be the affected neuronal physiology
responsible for the change in rotational behavior.
Key words:
genetic intervention; herpes simplex virus type 1 vectors; protein kinase C; nigrostriatal neurons; motor behavior; basal
ganglia
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INTRODUCTION |
The alterations in synaptic
transmission that cause behavioral adaptations are thought to be
mediated by changes in specific biochemical pathways in particular
types of neurons (Hebb, 1949 ), but the complexity of brain anatomy and
physiology has complicated experimental demonstrations. In
invertebrates, signal transduction pathways mediate behavioral
adaptations, including the gill withdrawal reflex in Aplysia
and long-term associative learning in Drosophila (Dudai,
1989). In contrast, in mammals, these pathways mediate short-term
changes in neuronal physiology (Kazmarek and Levitan, 1987), but
relatively little is known about the capability of a specific signal
transduction pathway, acting in a particular type of neuron, to direct
a change in behavior.
The protein kinase C (PKC) pathway is a promising candidate for
mediating changes in synaptic transmission that direct changes in
behavior (Tanaka and Nishizuka, 1994). PKC substrates play critical
roles in neuronal physiology, and activation of PKC, usually by phorbol
esters, can increase the release of various classical neurotransmitters
from diverse neuronal systems (Pozzan et al., 1984; Nichols et al.,
1987; Coffey et al., 1993). Nonetheless, evidence for the involvement
of PKC in specific behaviors remains indirect: PKC may be activated in
specific neurons in response to pain (Mao et al., 1992) or during
lordosis (Kow et al., 1994). During long-term potentiation (LTP),
hippocampal neurons contain a constitutively active catalytic domain of
PKC (Sacktor et al., 1993) that resembles the catalytic domains of
PKC produced by calpain (Kishimoto et al., 1989). PKC knock-out mice
display defective regulation of LTP (Abeliovich et al., 1993a) and mild deficits in selected learning paradigms (Abeliovich et al., 1993b). However, concomitant defects in cerebellar development and motor coordination (Chen et al., 1995; Kano et al., 1995), as well as the
absence of PKC from every cell in the mouse, complicate the interpretation of these experiments.
The nigrostriatal dopamine (DA) system is one of the few systems in
which a significant body of evidence suggests that the function of a
specific type of neuron regulates a specific set of behaviors:
degeneration of substantia nigra pars compacta (SNc) neurons underlies
Parkinson's disease; ablation (Ungerstedt, 1971) or electrical
stimulation (Arbuthnott and Ungerstedt, 1975) of SNc neurons can alter
rat motor behavior; SNc neuron activity (Diana et al., 1989) and
striatal DA release (Yamamoto et al., 1982) are increased in rats
trained to circle; and rats, cats, and humans exhibit a low level of
spontaneous rotational behavior, probably because of hemispheric
imbalances in the nigrostriatal system (Jerussi and Glick, 1976; Glick
et al., 1981; Gospe et al., 1990).
The capability of the PKC pathway in SNc neurons to modulate rotational
behavior was examined via a three-part genetic intervention strategy
(Geller et al., 1991): (1) microinjection of a Herpes simplex virus
type 1 (HSV-1) vector restricts gene transfer to a specific brain area
(During et al., 1994); (2) a cell type-specific promoter, such as a
preproenkephalin promoter (Kaplitt et al., 1994) or a tyrosine
hydroxylase (TH) promoter (Song et al., 1997), targets expression to a
specific type of neuron; and (3) a constitutively active signal
transduction enzyme, such as an adenylate cyclase, causes long-lasting
activation of a specific signaling pathway, thereby altering neuronal
physiology, such as increasing neurotransmitter release (Geller et al.,
1993). We used this genetic intervention strategy to introduce a
constitutively active fragment of rat PKCII (Pkc, encoded by
pkc) into SNc neurons. In cultured neurons, Pkc caused a
long-lasting, activation-dependent increase in neurotransmitter release. In the brain, microinjection of HSV-1 vectors that contain the
TH promoter targeted expression to SNc neurons. Expression of pkc in
a small percentage of the SNc neurons resulted in a long-term change in
apomorphine-induced rotational behavior. The magnitude of the change
was correlated with the number of affected SNc neurons, and
D2 DA receptor density was increased in the regions of the
striatum innervated by these neurons. Thus, this modulation of
apomorphine-induced rotational behavior was produced by alterations in
dopaminergic neurotransmission, arising from activation of the PKC
pathway in SNc neurons.
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MATERIALS AND METHODS |
Vectors. pHSVlac (Geller and Breakefield, 1988) and
pTHlac (Song et al., 1997) have been described. pTHlac contains a 6.8 kb fragment of the rat TH promoter (Brown et al., 1987). Constructions were performed by standard recombinant DNA procedures (Maniatis et al.,
1982). PKCII (PKC-II; Knopf et al., 1986) was partially digested
with an enzyme that yields a blunt end [nucleotides 719, 749, 831, 871, and 994 (pkc)], and the appropriate ClaI linker was
ligated (5'-CATCGATG-3' or 5'-CCATCGATGG-3' or 5'-CCCATCGATGGG-3'; New
England Biolabs, Beverly, MA) for fusion to the 3' end of flag
(Prickett et al., 1989) in pHSVflag (Geller et al., 1993); the 3' end
was an EcoRI site 3' to the cDNA. In pkc, 3' untranslated sequences were deleted to nucleotide 2166, just beyond the termination codon (nucleotides 2160-2162). The sequence surrounding the ATG (5'-TAAGCTTACCATGG-3') contains a Kozak consensus translation start
sequence and both HindIII and NcoI sites that
were used in subsequent constructions (see below). So that an in
situ hybridization probe could distinguish pkc transcription
unit RNA from endogenous rat PKC mRNAs, all of the pkc vectors
used in this study (for expression in mammalian cells) contain a 314 bp
fragment from the 3' end of the lacZ gene, and this fragment
is located in the 3' untranslated region of the vectors just before the
SV40 early region polyadenylation site (Geller and Breakefield, 1988;
Geller et al., 1993).
We isolated two point mutations in pkc to enable experimental
demonstrations that the effects of Pkc on neuronal physiology are
attributable to PKC enzyme activity. An absolutely conserved Lys
residue, present in subdomain II of all members of the protein kinase
family, is required for efficient phosphoryl transfer (Hanks et al.,
1988). Mutation of this Lys residue is a standard method for
inactivating protein kinases (for review, see Hanks et al., 1988),
including PKC (Ohno et al., 1990), and the corresponding Lys codon in
pkc was replaced with either a Gly or an Arg codon. Two fragments
that contain the mutation (generates a BamHI site) and
extend 5' or 3' were produced by PCR [template, pHSVpkc; primers, 5' fragment, 5'-CTACAAAGACGATGACGATAAATCG-3' (from flag; Prickett et al., 1989) and 5'-CCACATCTTTCTTCAGGATCC(C, Gly;
G, Arg)CACGGC-3' (complementary to PKCII nucleotides 1245-1272
[Knopf et al., 1986] except for the mutation); 3' fragment,
5'-GCCGTG(G, Gly; C, Arg)GGATCCTGAAGAAAGATGTGG-3' (PKCII nucleotides
1245-1272 except for the mutation); and
5'-GATCTACTTAGCTCTTGACTTCGGG-3' (complementary to PKCII
nucleotides 2145-2169)]. After digestion (5' fragment,
KpnI and BamHI; 3' fragment, BamHI and
BstBI), fragments were inserted into pHSVpkc
(KpnI and BstBI) to yield pHSVpkcGG and
pHSVpkcAA.
YEp51pkc, YEp51pkcAA, and YEp51pkcGG were constructed with the
YEp51 vector, previously used to express mammalian PKC genes in
Saccharomyces cerevisiae (Riedel et al., 1993). YEp51 was
digested with HindIII and BamHI; each of the
three pHSVpkc vectors (pHSVpkc, pHSVpkcAA, and pHSVpkcGG) was
digested partially with HindIII and to completion with
BglII; and the fragments that contain pkc, pkcAA, or
pkcGG were inserted into YEp51.
pTHpkc and pTHpkcGG were designed to express an mRNA essentially
identical to that expressed by pHSVpkc except for the addition of a
short sequence (<30 nucleotides) immediately after the
transcription start site of the TH promoter. The lacZ
gene in pTHlac (Song et al., 1997) was replaced with the intron
following the HSV immediate early (IE) 4/5 promoter [isolated by PCR:
template, pHSVpkc (Geller et al., 1993); primers,
5'-GGGAAGCTTACGGCGCCGGCCACGAACGACGGG-3' and
5'-GCCATGGTGCTTATCGACGAGGACGTTCTTCC-3'; PCR products
were digested with HindIII and NcoI] and either
pkc or pkcGG (NcoI and BglII)]. To verify
gene expression from these vectors, we infected catecholaminergic cell
lines; 1 d later both pkc transcription unit RNAs and flag-IR
positive cells were detected.
Packaging vectors into HSV-1 particles. CV1 monkey
fibroblasts were maintained in DMEM supplemented with 10% fetal bovine serum (FBS). M64A cells were maintained in DMEM supplemented with 5%
FBS and hypoxanthine aminopterin thymidine (HAT) medium, and E5 cells
were maintained in DMEM supplemented with 10% FBS and 0.5 mg/ml G418.
Initial candidate vectors and vectors for DA release experiments were
packaged (Geller et al., 1990) with HSV-1 strain 17 D30EBA (partial
deletion of IE 3; Patterson and Everett, 1990) and M64A cells (Davidson
and Stow, 1985). For all other experiments, including
3H-norepinephrine (NE) release and all in vivo
experiments, KOS strain d120 (complete deletion of IE 3) and E5 cells
(DeLuca et al., 1985) were used for packaging (Lim et al., 1996). The
titers [Geller et al. (1993); Song et al. (1997) and
immunocytochemistry section] of these vector stocks [which did not
contain detectable levels of wild-type HSV-1 (<10 pfu/ml)] were
2.0 × 106 infectious vector particles (IVP)/ml
pTHpkc and 2.6 × 107 plaque forming units
(pfu)/ml d120; 2.6 × 106 IVP/ml pTHpkcGG
and 1.4 × 107 pfu/ml d120; 1.4 × 107 IVP/ml pHSVpkc and 4.1 × 107 pfu/ml d120; 1.4 × 106
IVP/ml pTHlac and 1.6 × 107 pfu/ml d120; and
2.5 × 107 IVP/ml pHSVlac and 6.1 × 107 pfu/ml d120.
Neural cell culture and gene transfer. PC12 cells (Greene
and Tischler, 1976) were maintained in DMEM supplemented with 5% FBS
and 10% horse serum. Cultures of dissociated superior cervical ganglia
(Hawrot and Patterson, 1979) were prepared from 4-d-old Sprague Dawley
rats. [All work with rats was approved by Children's Hospital
Institutional Animal Care and Use Committee (IACUC).] Cultures were
treated with cytosine arabinoside (40 µM) on days 5-6
and used for expression studies within 2 weeks thereafter. PC12 cells
(5 × 105 cells/0.5 ml) or sympathetic cells
(~2 × 105 cells/0.5 ml) were infected with
vector stocks (~0.1 multiplicity of infection). Assays were performed
or cell extracts were prepared at 1 d after infection.
Antibodies. Polyclonal rabbit anti-flag antibody was raised
against NH2-MDYKDDDDKSC-NH2 coupled to bovine
serum albumin and affinity-purified by chromatography on SulfoLink
resin (Pierce, Rockford, IL) to which the peptide was coupled. Mouse
monoclonal anti-flag antibody (M-5) was kindly provided by Dr. M. Leahy
(Immunex, Seattle, WA). Affinity-purified rabbit anti-rat PKCII
antibody was obtained from Santa Cruz Biotechnology (Santa Cruz, CA).
Swine anti-rabbit immunoglobulin (Ig) G antibody and rabbit anti-HSV-1 particle antibody were obtained from Dako (Carpinteria, CA). Alkaline phosphatase-conjugated goat anti-mouse IgG antibody, alkaline phosphatase-conjugated goat anti-rabbit IgG antibody, mouse monoclonal anti-TH antibody, and alkaline phosphatase-conjugated rabbit
anti-digoxigenin antibody were purchased from Boehringer Mannheim
(Indianapolis, IN). Biotinylated goat anti-mouse IgG antibody and
biotinylated goat anti-rabbit IgG antibody were obtained from Jackson
ImmunoResearch Laboratories (West Grove, PA).
Immunocytochemistry. The cells were fixed with 4%
paraformaldehyde in PBS, pH 7.4, rinsed with PBS (three times for 5 min each), and immunocytochemistry was performed. The cells were incubated (overnight at 4°C) in PBS, 2% normal goat serum, 0.2% Triton X-100, and either mouse monoclonal anti-flag antibody (1:5000 dilution) or
rabbit anti-flag antibody (0.06 µg/ml). The cells were rinsed with
PBS, 0.2% Triton X-100 (three times for 10 min each), and then
incubated (2 hr, room temperature) with either alkaline
phosphatase-conjugated goat anti-mouse IgG antibody (1:2000 dilution)
or alkaline phosphatase-conjugated goat anti-rabbit IgG antibody
(1:2000 dilution) in PBS and 0.2% Triton X-100. The cells were rinsed
with 50 mM Tris-HCl, pH 7.4 (three times for 10 min each),
and alkaline phosphatase activity was visualized by the BCIP/nitroblue
tetrazolium (NBT) substrate (Sigma, St. Louis MO) in the presence of
levamisole (Sigma) to inhibit endogenous alkaline phosphatase activity.
Flag-IR positive cells were counted under 20× magnification.
Expression of recombinant PKCs in yeast. YEp51pkc,
YEp51pkcAA, and YEp51pkcGG use the S. cerevisiae Gal
10 promoter to regulate expression of the recombinant PKCs, and
these three plasmids were introduced into S. cerevisiae 334 (Hovland et al., 1989). S. cerevisiae transformants were
grown to 0.5 optical density (600 nm) at 30°C in minimal medium
lacking Leu and containing 2% D-glucose. Expression of
recombinant PKCs was induced by the addition of 0.1 vol of 20%
D-galactose. The cells were harvested by centrifugation
after an additional 2 hr of incubation, which resulted in levels of
recombinant PKC protein ranging from 0.2 to 1.0 ng/µg total cellular
protein.
Western blots. S. cerevisiae extracts and PC12
cell extracts were prepared by sonication and heating in 5 mM Tris-HCl, pH 8.0, 2 mM EDTA, and 2% SDS.
Soluble protein was collected after centrifugation for 10 min at
9000 × g. S. cerevisiae extracts, PC12 cell
extracts, and purified rat brain PKC [kindly provided by Dr. M. D. Browning (University of Colorado, Denver)] were subjected to
SDS-polyacrylamide gel electrophoresis and transferred
electrophoretically to nitrocellulose (0.2 µm; Costar, Cambridge,
MA). Protein transfer was recorded xerographically after Ponceau S
staining, and the blots were quenched/destained with
polyvinylpyrrolidone (Haycock, 1993). Then replicate blots were
incubated sequentially with primary antibody (1 µg/ml of either
rabbit anti-flag antibody or rabbit anti-rat PKCII antibody for 1 hr), secondary antibody (0.5 µg/ml of swine anti-rabbit IgG antibody
for 1 hr), and 125I-protein A (4 × 105 cpm/ml for 1 hr; New England Nuclear, Boston,
MA). Immunoreactivity was visualized by autoradiography.
PKC enzyme activity assays. Peptide substrates were
synthesized by the Louisiana State University (New Orleans, LA) Core
Labs, purified by reverse-phase HPLC with a C18 column (Vydac,
Hesperia, CA), and analyzed by electrospray mass spectrometry. Pellets
of S. cerevisiae were lysed by vortexing in an equal volume
of lysis buffer (50 mM Tris-HCl, 1 mM EGTA, 1 mM benzamidine, 20 µM leupeptin, and 0.1%
NP40, pH 8.0, at 0-4°C) and 2 vol of glass beads. Soluble protein
was collected after centrifugation for 10 min at 9000 × g. The disruption of the S. cerevisiae was
repeated twice, and the supernatants were combined and filtered (1.0 µm glass microfiber). A constitutively active fragment of purified
rat brain PKC was derived by partial digestion with trypsin (kindly
provided by Dr. Browning).
Aliquots (5 µl) of the S. cerevisiae extracts or the
proteolytically activated rat brain PKC (in extraction buffer) were
incubated with peptide substrates in a reaction mixture containing the
following (final concentrations in mM): 50 HEPES (pH 7.4, using Tris base), 10 magnesium acetate, 1 DTT, and 0.1 [-32P] ATP (500-1000 cpm/pmol; New England Nuclear)
plus 5 µM leupeptin and 50 nM okadaic acid.
Reactions (50 µl final volume) were initiated by the addition of the
ATP, incubated for 10 min at 30°C, and terminated by spotting on
Whatman P81 paper. Endogenous calcium-dependent kinase activity (e.g.,
S. cerevisiae PKC) was inhibited by the EGTA present in the
extraction buffer. The unincorporated 32P was removed by
rinsing the P81 paper in phosphoric acid, and the amount of
32P incorporation was determined by liquid scintillation
spectrometry (Roskoski, 1983). Relative 32P incorporation
values were calculated after subtracting the amount of 32P
incorporated in the absence of added substrate.
Neurotransmitter release assays. Cultured sympathetic
neurons from 2- to 4-d-old rats were used to study catecholamine
release. The release of endogenous DA, which affords greater detection sensitivity as compared with NE, was analyzed according to Geller et
al. (1993). One day after infection, the culture medium was replaced
with 0.2 ml of release buffer. Release buffer consisted of (in
mM) 135 NaCl, 3 KCl, 1 MgCl2, 1.2 CaCl2, 2 NaPO4, pH 7.4, and 10 glucose. "High K+" release buffer contained 56 mM KCl (replacing NaCl), and "Ca-free" release buffer
contained 0.1 mM EGTA in place of the CaCl2.
Release buffer was collected after a 15 min incubation at 37°C, and
one-tenth volume each of 2 M HClO4 and 1%
Na2S2O5 was added on ice.
Catecholamines and metabolites (with dihydroxybenzoic acid added as
internal standard) were separated by HPLC and analyzed with a serial
array of 16 electrode sensors (ESA, Chelmsford, MA). Peaks were
validated by comparison to standards (match criteria: retention
time ± 2%; peak width ± 3 sec; peak ratio between sensors
>80%). The amount of catecholamines in each sample was quantitated on
the basis of peak height (dominant sensor) relative to standards.
Efflux was calculated as pg DA/min per 106 cells,
and basal efflux refers to efflux in release buffer, whereas K+-dependent release refers to the difference
between basal efflux and efflux in high K+ release
buffer.
The release of previously accumulated [3H]-NE from
cultured sympathetic neurons was studied according to Wakade and Wakade (1988), using culture medium without serum supplement as the release buffer. One day after infection, cultures were rinsed and then incubated (1 hr at 37°C) in release buffer containing
[3H]-NE (0.1 µM, 15.4 Ci/mmol;
NET-377, New England Nuclear). After three 20 min rinses, cultures were
brought to room temperature, and 0.3 ml of release buffer containing 2 µM desipramine was added and collected every 3 min. After
four or five collections (at which point basal efflux had stabilized),
release was induced by switching to release buffer containing an
additional 30 mM KCl (with or without 1.2 mM
EGTA). At the end of the incubations, cells were lysed in 0.5 ml of
0.5% SDS. Radioactivity in the incubations and cell lysates was
determined by liquid scintillation spectroscopy, and efflux was
calculated as the percentage of the total radioactivity per well. Basal
efflux and K+-dependent release were calculated as
above.
As used within the context of K+-stimulated efflux
over a duration of 15 or 3 min, "release" is not intended to
represent either the initial rates of secretion or those processes
underlying the rapid kinetics of synaptic transmission.
Stereotactic injections and behavioral testing. Vector
stocks were purified and concentrated as described (Lim et al., 1996). Vector stocks were delivered by stereotactic injection into the midbrain [mediolateral (ML) 4.0, anteroposterior (AP) 3.5, and dorsoventral (DV) 6.8; ML 4.0, AP 3.7, and DV 6.8 (Paxinos and Watson,
1986); AP is relative to the interaural line, ML is relative to the
sagittal suture, and DV refers to the distance traveled from the dural
membrane with a 20° angle for the needle toward the midline] of male
Sprague Dawley rats (100-125 gm). These rats are smaller than those
used for the atlas (Paxinos and Watson, 1986), and the injection sites
were located just dorsal to the posterior SNc, as verified by cresyl
violet staining. Each rat received a total of 6 × 104 IVP (two sites and 3 µl/site for 6 µl of
total volume; 1 × 104 IVP/µl); each
injection was performed slowly and evenly over ~5 min.
We have shown previously (Song et al., 1997) that pTHlac targets
expression 10-fold to SNc neurons, as compared with pHSVlac, which uses
the HSV-1 IE 4/5 promoter. At 4 d after the injection of pTHlac we
observed ~950 X-gal positive cells per rat, ~40% SNc neurons
[assayed by double staining for either X-gal and TH-immunoreactivity (IR) or Escherichia coli -galactosidase-IR and TH-IR],
and with pHSVlac we observed ~1500 positive cells per rat, ~4% SNc
neurons (Song et al., 1997). We found (Song et al., 1997) that the
stability of expression (average positive cells per rat at 6 weeks
divided by average positive cells per rat at 4 d) was 44% for
pTHlac (not a statistically significant decrease) and 3% for pHSVlac.
Additionally, after the injection of pTHlac, no positive cells were
observed in other catecholaminergic areas (e.g., ventral tegmentum,
locus coeruleus) or in the striatum, and injection of pHSVlac resulted in very few positive striatal cells (Song et al., 1997). Localized gene
transfer with limited retrograde transport to distant neurons also was
observed after vectors were delivered into either the striatum (During
et al., 1994) or the hippocampus (Wood et al., 1994).
Rotational behavior was tested after the administration of apomorphine
(2.5 mg/kg, s.c.). In some tests, fluphenazine (0.2 mg/kg, s.c.) was
administered 3 hr before the apomorphine (Bruno et al., 1985). The net
rotations performed during each 5 min period for 1 hr after apomorphine
administration were measured by a computer-controlled rotameter
(Omnitech Electronics, Columbus, OH; Ungerstedt and Arbuthnott, 1970).
The net rotations performed during each 5 min period by each rat in a
group were used to calculate the averages and totals shown in Figures 2
and 4 and in Table 4. Behavioral data were analyzed by ANOVA and
Newman-Keuls tests.
Immunohistochemistry. The rats were anesthetized with
chloral hydrate (400 mg/kg, i.p.) and perfused with 50 ml of PBS, pH 7.4, followed by 200 ml of 4% paraformaldehyde in PBS. The brains were
cryoprotected, coronal sections (25 µm) were cut on a freezing microtome, and immunohistochemistry was performed on free-floating sections. Sections were preincubated in PBS and 0.3%
H2O2 (for 10 min at room temperature) and then
rinsed with PBS (three times for 5 min each). Sections were
permeabilized by incubation for 30 min at 37°C in PBS, 2% normal
goat serum, and 0.2% Triton X-100 and then incubated overnight at
4°C in the same buffer with the primary antibody [mouse monoclonal
anti-TH antibody (1:200 dilution), rabbit anti-flag antibody (0.15 µg/ml), or rabbit anti-HSV-1 particle antibody (1:2000 dilution)].
The sections were rinsed at room temperature with PBS and 0.2% Triton
X-100 (three times for 10 min each) and then were incubated for 2 hr at
room temperature in the same buffer with the secondary antibody
[either biotinylated goat anti-mouse IgG antibody or biotinylated goat
anti-rabbit IgG antibody (1:2000 dilution)]. The sections were rinsed
at room temperature with PBS and 0.2% Triton X-100 (three times for 10 min each), incubated (at room temperature for 1 hr) with
avidin-biotinylated peroxidase complex (ABC) reagent (Vector
Laboratories, Burlingame, CA), and rinsed. Immunoreactivity was
visualized with diaminobenzidine according to the manufacturer's
instructions.
With most of the rats, every eighth section was analyzed for TH-IR,
every fourth section was analyzed for flag-IR positive cells, and ~80
sections contained the SNc. The number of flag-IR positive SNc cells in
each of 11 rats in the pTHpkc group was determined by analyzing
every section developed for flag-IR that contained the SNc. In some
rats, HSV-1 particle-IR was analyzed in every eighth section. Positive
cells were counted by locating the cell under 5 or 10× magnification
and then verifying cellular morphology under 20 or 40×
magnification.
Detection of recombinant RNAs. For RT-PCR, brain regions
were dissected and rapidly frozen, and RNA was isolated (Ivins et al.,
1993). To remove any contaminating DNA, we centrifuged the RNA through
a CsCl gradient, we precipitated the resuspended pellet with a one-half
volume of ethanol (selectively precipitates RNA), and we treated the
remaining material with DNase (Ivins et al., 1993). Reverse
transcriptase was used to transcribe 1 µg of the RNA into cDNA, which
served as the template for PCR (Ivins et al., 1993). The conditions for
the PCR were 39 cycles of 94°C for 2 min, of 60°C for 3 min, and of
72°C for 3 min, with a final extension time of 10 min at 72°C. The
primer for reverse transcription, which also was used for PCR, was
complementary to vector sequences in the 3' untranslated region of the
pkc transcription unit (5'-TGACACCAGACCACTGGTAATGGT-3'), and the
other primer for PCR was from pkc [5'-AATGTGCCGGTGCCGCCGGAAG-3'; nucleotides 999-1020 of the rat PKCII cDNA (Knopf et al., 1986)]. The PCR products were subjected to Southern analysis, using a radiolabeled oligonucleotide from pkc
(5'-ACAATGGCAACAGGGACCGGATGAAACTGA-3'; nucleotides 1129-1158 of the
rat PKCII cDNA).
For in situ hybridization, the rats were perfused with 4%
paraformaldehyde (see Immunohistochemistry, above), and coronal brain
sections (20 µm) were cut on a freezing microtome. Sections on glass
slides were post-fixed with 4% paraformaldehyde, followed by graded
concentrations of ethanol. The hybridization probe was complementary to
lacZ nucleotides 3016-3079 (Kalnins et al., 1983) present
in the 3' untranslated region of the vectors (see Vectors, above) and
was synthesized by in vitro transcription [T3 RNA
polymerase, digoxigenin-conjugated UTP (Boehringer Mannheim)] of PCR
products [template, pHSVlac; 5' primer,
5'-AAAAAGAATTCCAGCTGAGCGCCGG-3' (5 A and lacZ nucleotides
3016-3035); 3' primer,
5'-AATTAACCCTCACTAAAGGGAAGAAATACGGGCAGACATGGCC-3' (a T3 polymerase
promoter [Sommer et al., 1990] and 20 nucleotides complementary to
SV40 early region polyadenylation sequences)]. After in
situ hybridization (Grant et al., 1993), the signal was visualized
with an alkaline phosphatase-conjugated rabbit anti-digoxigenin antibody (1:500 dilution) and the BCIP/NBT substrate (Boehringer Mannheim) with levamisole (Vector Laboratories). Sections were counterstained with methyl green and coverslipped with aqueous mounting
medium.
Detection of vector DNAs. Sections adjacent to those used
for immunohistochemistry were analyzed. The midbrain was extracted (0.5 mg tissue/µl) in proteinase K (1 mg/ml) buffer (Higuchi, 1989).
Samples were subjected to nested PCR, using primer pairs that recognize
the pkc transcription unit, but not the rat PKC gene. The primers
for the first reaction were 5'-AGGAGGAACGTCCTCGTCGATAAGC-3' [20
nucleotides from the HSV IE 4/5 intron (HSV-1 nucleotides 132,539-132,558 [McGeoch et al., 1988]), followed by five
nucleotides from the sequence surrounding the ATG (see Vectors,
above)] and 5'-TGGTTTATAAGGTGGCTGAATCTCC-3' (complementary to
nucleotides 1973-1997 of the rat PKCII cDNA). The primers for the
second reaction were 5'-CTACAAAGACGATGACGATAAATCG-3' (from the flag
sequence) and 5'-TTCGAGTTTCTCCCAGTCGATATACC-3' (complementary to
nucleotides 1942-1967 of the rat PKCII cDNA). The conditions for
both reactions were 40 cycles of 94°C for 1 min, of 55°C for 1 min,
and of 72°C for 4 min. The reaction products were displayed on a
1.2% agarose gel.
DA receptor radioligand-binding assays. After rapid
decapitation, the brains were removed quickly, frozen in isopentane at  15 to 20°C for ~2 min, and stored at 20°C. Sections (20 µm) were cut on a cryostat, thaw-mounted onto gelatin-coated slides, desiccated, and stored at 20°C. Sections were preincubated in TBSI
[containing (in mM) 50 Tris-HCl, pH 7.4, 120 NaCl, 5 KCl, 2 CaCl2, and 1 MgCl2] at room
temperature for 5 min. To quantify the number of receptors of the
D2 subfamily, we incubated sections at room temperature for
60 min in TBSI containing 0.7 nM
[3H]spiperone (123.0 Ci/mmol; Amersham, Arlington
Heights, IL) and 40 nM ketanserin (Janssen Pharmaceuticals,
Titusville, NJ) to block radioligand binding to 5-HT2 receptors. After
incubation, the slides were rinsed (two washes for 20 sec each) in
ice-cold TBSIA (TBSI plus 0.02% ascorbic acid) and then in ice-cold
H20 (one wash for 20 sec) and rapidly dried by aspiration
and mild heating. Nonspecific [3H]spiperidol
binding, defined with 1 µM (+)butaclamol (Research Biochemicals, Natick, MA), constituted ~25% of the total binding. To
quantify the number of receptors of the D1 subfamily, we
preincubated sections as above, incubated them for 60 min in TBSIA
containing 1.0 nM [3H]SCH23390 (87.0 Ci/mmol; New England Nuclear), and rinsed and dried them as described
above. Nonspecific binding, defined with 5 µM
(+)butaclamol, was ~10% of the total binding.
To quantitate DA receptor densities, we apposed slides and
[3H]-containing plastic standards (American
Radiolabeled Chemicals, St. Louis, MO) to
[3H]-sensitive Hyperfilm (Amersham) for 12-14 d
at 20°C and developed the film with Kodak (New Haven, CT) D19
developer. Autoradiographic images were analyzed by computer-assisted
densitometry (MCID; Image Research, St. Catherine's, Ontario, Canada).
Density was linearized by calibration against the appropriate
radiolabeled plastic standards. The contribution of nonspecific binding
was removed from the images of total binding, using a pixel-by-pixel subtraction routine, to yield images of specific binding. Binding density was quantified from these specific binding images, and readings
were taken from the four quadrants of the anterior, medial, and
posterior caudate putamen in each hemisphere. The binding data were
analyzed by ANOVA, followed by post hoc comparisons.
| |
RESULTS |
Mutations in PKC
To enable genetic activation of PKC pathways in neurons, we
isolated a constitutively active mutant of PKC. Rat PKCII [673 amino acids (aa)] contains a regulatory domain (aa 1-339) and a
catalytic domain (aa 340-603) (Knopf et al., 1986; Hanks et al.,
1988). Five candidate deletions of the PKCII cDNA, encoding proteins
that extend from aa 194, 204, 231, 245, or 285 to the C terminus of
PKCII, were isolated. To allow for the detection of these
recombinant PKCs in the presence of endogenous PKCs, we fused the 5'
ends of these candidate deletions to 10 codons that encode the flag
epitope tag (Prickett et al., 1989). PC12 cells were infected with
HSV-1 vectors designed to express each of the candidate deletions, and
1 d later flag-IR positive cells were visualized. Expression of
the deletion that encodes aa 285 to the C terminus of PKCII resulted
in the highest level of flag-IR (data not shown); the gene and protein
were designated pkc and Pkc, respectively, and pkc was used in
the subsequent experiments.
As shown in Figure 1, Pkc and PkcGG
(a Lys-to-Gly point mutation designed to lack enzyme activity; see
Vectors above) were detected readily in PC12 cells infected with either
pHSVpkc or pHSVpkcGG. A Mr ~50,000 band
(calculated molecular weight 45,800) was detected by Western blots with
either affinity-purified rabbit anti-flag antibody or anti-PKCII
antibody (Fig. 1A), and flag-IR positive cells were
detected by immunocytochemical analysis with either a rabbit anti-flag
antibody (Fig. 1B,C) or a mouse monoclonal anti-flag
antibody (data not shown). PkcAA (a Lys-to-Arg point mutation) also
was detected by using these methods (data not shown).
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Figure 1.
Western blots and immunocytochemical analyses of
Pkc, PkcGG, and PkcAA in S. cerevisiae and PC12
cells. A, Replicate sets of aliquots of SDS-solubilized
S. cerevisiae and PC12 cells were subjected to
SDS-polyacrylamide gel electrophoresis, followed by blot immunolabeling
analysis. Protein staining is shown in the left panel
(PONCEAU), and immunoreactivities with the
indicated antibodies are shown in the right panels.
Lane 1, Purified rat brain PKC; lanes
2-5, extracts from S. cerevisiae harboring
no YEp51 plasmid, YEp51pkc, YEp51pkcAA, or YEp51pkcGG,
respectively; lanes 6-8, extracts from PC12 cells
1 d after infection with pHSVlac, pHSVpkc, or pHSVpkcGG,
respectively. The molecular weight markers are -gal,
-galactosidase (116 kDa); phos, phosphorylase b (97 kDa); BSA, bovine serum albumin (67 kDa);
cat, catalase (60 kDa); GDH, glutamate
dehydrogenase (55 kDa); oval, ovalbumin (42 kDa); and
CA, carbonic anhydrase (29 kDa). B, C,
Flag-IR positive cells 1 d after PC12 cells were infected with
either pHSVpkc (B) or pHSVpkcGG
(C). Scale bars, 333 µm.
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|
Pkc exhibits a substrate specificity similar to rat
brain PKC
As shown in Figure 1A, pkc, pkcAA, and
pkcGG were expressed at relatively comparable levels in the yeast
S. cerevisiae, using the YEp51 vector (Riedel et al., 1993).
As measured by using a peptide substrate for PKC derived from the
MARCKS protein (Stumpo et al., 1989), PKC activity was detected only in
the extracts of S. cerevisiae that harbored YEp51pkc and
not in extracts from S. cerevisiae that harbored either
YEp51pkcAA or YEp51pkcGG (Table 1).
Also, as shown in Table 2, the relative
selectivity of protein kinase activity in the extracts of S. cerevisiae expressing pkc was comparable to that exhibited by
purified catalytic fragment of rat brain PKC (made constitutively
active by partial digestion with trypsin).
Pkc causes a long-lasting, activation-dependent increase in
catecholamine release
To determine whether Pkc could alter neuronal physiology, we
measured neurotransmitter release from cultured sympathetic cells (1 d
after infection) as the increase in efflux of either endogenous DA or
previously accumulated [3H]-NE produced by
elevated (depolarizing) levels of K+ in the release
buffer. As shown in Table 3, elevated
K+ (56 mM) produced substantially more
DA release from cells infected with pHSVpkc, and this effect was
dependent on extracellular Ca2+ (data not shown).
However, neither endogenous NE nor endogenous DA under the other
conditions could be quantitated (for lack of sensitivity). Subsequent
experiments examined the release of [3H]-NE from
cultures in which endogenous pools were prelabeled via high-affinity
uptake. As shown in Table 3, previous infection with pHSVpkc
resulted in a modest but reliable increase in basal efflux and a
larger, 35-50% increase in K+-dependent release
(30 mM K+). The
K+-dependent release of [3H]-NE
was calcium-dependent in each condition (data not shown). In separate
cultures, flag-IR was detected in cells that displayed either
neurofilament-IR or glial morphology (data not shown); however, a glial
provenance of DA and NE seems unlikely because glia do not take up,
store, or release significant amounts of catecholamines (Hansson,
1983). Additionally, the flag-IR in cells displaying a neuronal
phenotype usually was restricted to the cell bodies and proximal
processes.
Expression of pkc in SNc neurons causes changes in
apomorphine-induced rotational behavior
Our hypothesis was that expression of pkc in SNc neurons might
alter the physiology of the nigrostriatal DA system, with consequent
effects on motor behavior. Because Pkc produced a long-lasting,
activation-dependent increase in catecholamine release from cultured
neurons, we reasoned that physiological activation of SNc neurons that
contain Pkc might result in increased DA release from the striatal
varicosities/terminals of the affected neurons. Changes in striatal DA
levels are known to alter striatal DA receptor levels and/or
sensitivity, and such changes in the levels/sensitivity of DA receptors
unilaterally can be revealed by treatment with a DA receptor agonist,
such as apomorphine, resulting in asymmetrical rotational behavior
(Ungerstedt and Arbuthnott, 1970; Creese and Snyder, 1977; Neve et al.,
1984). Unilateral lesions of SNc neurons result in decreased striatal DA levels, increased striatal DA receptors levels, and rotations in the
contralateral direction. Thus, the lesion model predicts that
expression of pkc in SNc neurons unilaterally would increase striatal DA release, decrease the levels/sensitivity of striatal DA
receptors, and result in apomorphine-induced rotational behavior in the
ipsilateral direction. However, other predictions can be supported. For
instance, some investigators have reported that drugs that increase DA
release also increase striatal DA receptor levels (Klawans et al.,
1979; Wilner et al., 1980). Furthermore, spontaneous rotational
behavior is associated with specific and distinct changes in striatal
DA receptor levels (Jerussi and Glick, 1976; Glick et al., 1981). In
light of these contradictory results concerning changes in striatal DA
receptor levels, we predicted that expression of pkc in SNc neurons
unilaterally would increase striatal DA release, cause a change (either
an increase or a decrease) in the levels of striatal DA receptors, and
cause apomorphine-induced rotational behavior in the ipsilateral
direction.
This hypothesis was tested by using the TH promoter to target the
expression of pkc to SNc neurons and then measuring the effect on
apomorphine-induced rotational behavior. On the basis of previous
results (Song et al., 1997) (see Stereotactic Injections and Behavioral
Testing above), which demonstrated that midbrain injections of a HSV-1
vector containing the TH promoter (pTHlac) target expression to SNc
neurons (10-fold as compared with pHSVlac, which contains the HSV IE
4/5 promoter), vectors expressing either pkc or pkcGG from the TH
promoter (pTHpkc and pTHpkcGG, respectively) were isolated.
Before gene transfer, most rats exhibited minimal apomorphine-induced
rotational behavior (Fig.
2A, Table
4). From 5 to 7 d later, pTHpkc,
pTHpkcGG, pHSVpkc, pTHlac, or vehicle alone (PBS) was injected
into the midbrain of these rats. Apomorphine-induced rotational
behavior was measured at 1, 2, and 3 weeks after gene transfer. The
rats in the pTHpkc group displayed increases in both the maximum
rotation rate (Fig. 2) and the total number of rotations (Table 4),
which were in the direction ipsilateral to the microinjection. The
increase in the number of rotations was statistically significant when
compared with either the number of rotations before gene transfer or
the number of rotations exhibited by the rats in the control groups
(Table 4). In contrast, rats in the control groups (pTHpkcGG,
pHSVpkc, pTHlac, or PBS) displayed no statistically significant
changes in rotational behavior (Fig. 2B, Table 4),
demonstrating that the change requires both enzymatically active Pkc
and its targeted expression to SNc neurons. At week 4 after gene
transfer, the administration of fluphenazine, a DA receptor antagonist,
before apomorphine blocked the apomorphine-induced rotational behavior
(Fig. 2A, Table 4). Moreover, rotational asymmetry
was shown to persist by retesting the rats in the pTHpkc and pTHlac
groups with apomorphine only at week 5 (Table 4).
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Figure 2.
Apomorphine-induced rotational behavior after
microinjection of pTHpkc, control vectors, or PBS. The number of
rotations was measured during each 5 min period for 1 hr after
apomorphine administration. The rats were tested for
apomorphine-induced rotational behavior before gene transfer
(pretest). From 4 to 7 d later, pTHpkc,
pTHpkcGG, pHSVpkc, pTHlac, or PBS was injected into the midbrain.
Apomorphine-induced rotational behavior was measured at 1, 2, and 3 weeks after gene transfer. On week 4, fluphenazine, a DA receptor
antagonist, was administered 3 hr before apomorphine-induced rotational
behavior was tested (Bruno et al., 1985). On week 5, rats in the
pTHpkc and pTHlac groups were retested for apomorphine-induced
rotational behavior. A, For the rats in the pTHpkc
group, shown is the average number of rotations performed during each 5 min period in each of five tests (pretest and
weeks 1-4). These rotations were in the
ipsilateral direction. B, For the rats in the pTHpkc
and control groups, shown is the average over three tests (weeks
1-3) of the average number of rotations performed during each
5 min period.
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|
Recombinant gene expression in the midbrain
Flag-IR positive cells were detected at 4 d after gene
transfer as well as after completion of the behavioral analysis.
Low-power photomicrographs of adjacent sections from rats killed at
4 d after gene transfer with pTHpkc show a number of flag-IR
positive cells present in the SNc (Fig.
3B), as delineated by TH-IR
(Fig. 3A). A high-power view shows that the flag-IR is
located in the cell bodies and proximal processes of cells that display
a neuronal morphology (Fig. 3C). Flag-IR was assayed at 6 weeks after gene transfer with pTHpkc in some of the rats that had
undergone behavioral testing: a low-power photomicrograph shows ~12
flag-IR positive SNc cells (Fig. 3D) and a higher power view
shows a cell with flag-IR in the cell body (Fig. 3E). The
flag-IR positive SNc cells also had neuronal morphology (Fig.
3F-H). From 4 to 6 weeks after gene transfer with
pTHpkcGG, flag-IR positive SNc cells with neuronal morphology were
detected (data not shown). These results are similar to our previous
results (Song et al., 1997), which showed that, at 4-6 weeks after the
injection of pTHlac, both X-gal positive and -galactosidase-IR
positive SNc cells with neuronal morphology were observed. The level of
expression from the TH promoter declined somewhat over time with
pTHpkc. At 4 d after gene transfer the flag-IR usually extended
to the proximal processes (Fig. 3B,C), but by 4-6 weeks the
flag-IR frequently was restricted to the cell body (Fig.
3D-H), and we have reported an analogous decline in
the level of expression, using pTHlac (Song et al., 1997). The two
injection sites were located proximal to the posterior SNc, and after
injection of pTHlac, pTHpkc, or pTHpkcGG, the majority of the
cells that contained recombinant gene products was located in the
posterior region of the SNc (data not shown).
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Figure 3.
TH-IR positive cells and flag-IR positive cells
with neuronal morphology at either 4 d or 6 weeks after gene
transfer with pTHpkc. pTHpkc was injected into the midbrain;
either 4 d (A-C) or 6 weeks
(D-H) later (after the behavioral
analysis), adjacent sections were analyzed for TH-IR and flag-IR
positive cells. The SNc was located in sections by visualizing the
TH-IR positive cells (A). A low-power view of an
adjacent section (B) shows flag-IR positive cell
bodies and proximal processes in SNc, but not in SNr. Three flag-IR
positive cell bodies are indicated by arrows, and a
high-power view (C) shows that these
three cells display neuronal morphology, including neuronal-like
processes. D, A low-power view shows ~12 flag-IR
positive cells in the SNc at 6 weeks after gene transfer. Each
arrow indicates one to several flag-IR positive cells,
and the flag-IR positive cell indicated by the empty
arrow is shown under medium power in E.
F, High-power view of a flag-IR positive SNc cell with a
large cell body, characteristic of a SNc neuron, from a second rat.
G, H, Low- and high-power views of a flag-IR positive
SNc cell, with processes characteristic of a SNc neuron, from a third
rat. SNc, Substantia nigra pars compacta;
SNr, substantia nigra pars reticulata. Scale bars:
A and D, 80 µm; B, 45 µm; C, 25 µm; F, 20 µm;
G, 50 µm; H, 10 µm.
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The number of flag-IR positive cells, determined by cell counts,
represented a small percentage of the number of neurons in the SNc. At
4-6 weeks after gene transfer using pTHpkc, we observed ~20-100
flag-IR positive SNc cells, and we have reported previously that the
use of pTHlac resulted in ~100-500 X-gal positive SNc cells (Song et
al., 1997). A likely explanation for the approximately fivefold higher
number of X-gal positive cells than flag-IR positive cells is that the
flag-IR assay is not so sensitive as the X-gal assay. It is unlikely
that the pkc transcription unit RNA is highly unstable, because this
RNA was detected by both RT-PCR and in situ hybridization
(see next section). It is also unlikely that the constitutively active
PKC caused a downregulation of the TH promoter in the vector, because
similar numbers of flag-IR positive SNc cells were observed after the
injection of equal titers of either pTHpkc or pTHpkcGG.
Expression from pTHpkc was relatively stable, and approximately two-
to threefold more flag-IR positive cells were observed at 4 d as
compared with 4-6 weeks after gene transfer, similar to the results
previously obtained with pTHlac [approximately twofold more X-gal
positive cells at 4 d as compared with 6 weeks (Song et al.,
1997)]. Thus, it is probable that pTHpkc supported the expression
of pkc in ~0.1-2% (correcting for the sensitivity of the flag-IR
assay) of the ~22,000 neurons in the SNc (Poirier et al., 1983)
throughout the experimental period.
The relationship between the number of flag-IR positive cells and the
amount of rotational behavior was examined by using data from
individual rats in the pTHpkc group. The number of flag-IR positive
SNc cells was determined in each of 11 rats (killed at 4-6 weeks after
gene transfer), and the average of the total number of rotations for
weeks 1, 2, and 3 was calculated for each rat. Of note, the number of
flag-IR positive SNc cells displayed a statistically significant
correlation with the amount of rotational behavior (Fig.
4).
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Figure 4.
Apomorphine-induced rotational behavior as a
function of the number of flag-IR positive SNc cells. The average of
the total number of rotations performed during each of three tests
(weeks 1-3) is plotted for each of 11 rats that received pTHpkc as
a function of the number of flag-IR positive cells in each rat.
Least-squares regression analysis of these data (with and without
values from the rat indicated by the circled square)
yielded correlation coefficients of r = 0.74 (p < 0.01; solid line) and
r = 0.89 (p < 0.001;
dashed line), respectively.
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The location of the flag-IR positive cells was compared after the
injection of pTHpkc or pHSVpkc. At 4 d after gene transfer with pTHpkc, approximately one-half of the flag-IR positive cells were located in the SNc (see Fig. 3); in contrast, with pHSVpkc, only ~5% of flag-IR positive cells were located within the SNc (data
not shown). Similarly, we previously reported that at 4 d after
the injection of pTHlac or pHSVlac, respectively, 40 or 4% of the
X-gal positive cells contained TH-IR (Song et al., 1997). Other
differences in the expression of pkc from pTHpkc and pHSVpkc offer little more to explain why only pTHpkc altered rotational behavior; because expression from the IE 4/5 promoter is relatively unstable [pHSVlac, ~30-fold decrease in the number of positive cells
between 4 d and 6 weeks after gene transfer (Song et al., 1997)]
and because the flag-IR assay was not so sensitive as the X-gal assay,
few flag-IR positive cells were observed at 4-6 weeks after gene
transfer with pHSVpkc, although pkc transcription unit RNA was
still detected (see next section). Of note, analogous to results
obtained with pHSVlac and pTHlac (Song et al., 1997), more flag-IR
positive cells were observed with pHSVpkc than with pTHpkc at
4 d after gene transfer, but at 1 week after gene transfer only
pTHpkc directed rotational behavior (see Fig. 2, Table 4). Additionally, although pTHpkc and pHSVpkc may express different levels of pkc transcription unit RNA because of their different promoters, Pkc is constitutively active, and threshold levels sufficient to cause physiological effects are likely to be low. Thus,
expression of pkc must be targeted to SNc cells to cause rotational
behavior.
Detection of recombinant RNAs and persistence of vector DNAs
The long-term transcriptional activities of pTHpkc and
pHSVpkc were analyzed by using both RT-PCR and in situ
hybridization. At 2-6 weeks after gene transfer, pkc transcription
unit RNA was detected in the midbrain (using RT-PCR), but not in the
cerebellum, in each of three rats that received pTHpkc and in each
of three rats that received pHSVpkc (Fig.
5A). Such results are similar to those previously reported in which lacZ RNA was detected
in the midbrain, but not in the cerebellum, at 6 weeks after gene transfer with pTHlac (Song et al., 1997). In situ
hybridization was used to localize pkc transcription unit RNA. Two
weeks after the injection of pTHpkc, pkc transcription unit RNA
was detected in SNc cells from each of three rats that were analyzed
(Fig. 5B,C), but pkc RNA was not detected in SNc cells
from one rat that received only helper virus (d120; data not
shown).
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Figure 5.
Long-term expression of pkc transcription unit
RNA from pTHpkc and pHSVpkc and the persistence of vector DNAs.
A, Detection of pkc transcription unit RNA by RT-PCR.
At 2 weeks (pTHpkc) or 6 weeks
(pHSVpkc) after injection into the midbrain,
RNA was isolated from either the midbrain or the cerebellum. The RNA
was treated with DNase, and reverse transcription was performed by
using a primer from the 3' untranslated region of the pkc
transcription unit. The reverse transcription reaction products were
amplified by performing PCR with primers from the pkc transcription
unit. The PCR reaction products were detected by Southern blot
analysis, using a radiolabeled oligonucleotide from the pkc coding
region. The predicted size of the RT-PCR products is 1239 bp.
Mb, Midbrain; Cb, cerebellum. B,
C, Detection of pkc transcription unit RNA by in
situ hybridization. At 2 weeks after the injection of
pTHpkc, in situ hybridization was performed with a
digoxigenin-labeled probe. Hybridization was visualized by using an
alkaline phosphatase-conjugated anti-digoxigenin antibody, and the
nuclei were detected by counterstaining with methyl green.
B, A camera lucida drawing of a section that was
analyzed; the rectangle shows the location of a group of
positive cells in the SNc. C, A high-power
photomicrograph reveals a number of positive cells. Each of the two
arrows points to an area of hybridization signal
proximal to a nucleus. Scale bar, 50 µm. D-G,
Detection of vector DNAs by PCR. DNA was extracted from the midbrain in
sections adjacent to those used for immunohistochemistry, PCR was
performed with primers from the pkc transcription unit, and the PCR
reaction products were displayed on an agarose gel. The predicted size
of the PCR reaction products is 1003 bp. D, All of the
rats were analyzed at 4 d after gene transfer. Lanes
1, 2, pTHpkc or pHSVpkc DNA isolated from
E. coli, respectively; lanes 3-5, three
rats that received pHSVpkc; lanes 6-9, four rats
that received pTHpkc; lanes 10-12, three rats that
received PBS; lane 13, plasmid PKC-II DNA [contains the
rat PKCII cDNA (Knopf et al., 1986)]; lane 14, no
DNA; lanes 15, 16, no primers or no
Taq polymerase, respectively, with pTHpkc DNA
isolated from E. coli. E-G, All of the
rats were analyzed at 6 weeks after gene transfer. E, Lane
1, pTHpkc DNA isolated from E. coli;
lanes 2, 3, two rats that received
pHSVpkc; lanes 4-11, eight rats that received
pTHpkc; lane 12, one rat that received
PBS; lane 13, plasmid
PKC-II DNA; lanes 14, 15, no primers or
no Taq polymerase, respectively, with pTHpkc DNA
isolated from E. coli. F, Lanes 1,
2, pTHpkcGG or pTHpkc DNA isolated from E.
coli, respectively; lanes 3-6, four rats that
received pTHpkc (in addition to the eight rats in E);
lanes 7-9, three rats that received pTHpkcGG;
lane 10, one rat that received PBS. G, A
BamHI site is predicted to be present in the PCR
reaction products obtained by using pTHpkcGG, but not pTHpkc, as
the template; digestion with BamHI is expected to result
in 288 and 715 bp fragments. The first two nucleotides in the
BamHI site represent the last two nucleotides in the Gly
codon in pkcGG that replaced the evolutionarily conserved Lys codon
in pkc. Lanes 1, 2, pTHpkcGG DNA
isolated from E. coli, either undigested or digested
with BamHI, respectively; lanes 3-6, two
rats that received pTHpkc, either undigested (lanes
3, 5) or digested with BamHI
(lanes 4, 6); lanes
7-10, two rats that received pTHpkcGG, either undigested
(lanes 7, 9) or digested with
BamHI (lanes 8, 10).
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Persistence of vector DNAs was demonstrated by performing PCR on DNA
extracted from sections adjacent to those used for
immunohistochemistry. At 4 d after gene transfer, pkc
transcription unit DNA sequences were detected in four rats that
received pTHpkc and three rats that received pHSVpkc, but pkc
transcription unit DNA sequences were not detected in three rats that
received PBS (Fig. 5C). At 6 weeks after gene transfer,
pkc transcription unit DNA sequences were detected in 12 rats that
received pTHpkc, three rats that received pTHpkcGG, and two rats
that received pHSVpkc, but no signal was detected in two rats that
received PBS (Fig. 5D,E). The mutation in pTHpkcGG was
preserved: a BamHI site was created in constructing pkcGG
from pkc, and the last two nucleotides of the inserted Gly codon
represent the first two nucleotides of the BamHI site. The
PCR products produced from DNA of two rats that received pTHpkcGG
were cleaved by BamHI to yield two bands with the predicted
sizes, but the PCR products obtained from the DNA of two rats that
received pTHpkc were resistant to digestion by BamHI
(Fig. 5F). Similar results have been reported after
gene transfer with either pTHlac or pHSVlac (Song et al., 1997).
Gene transfer with pTHpkc results in changes in the striatal
DA system
Following the hypothesis that the change in apomorphine-induced
rotational behavior after gene transfer with pTHpkc resulted from an
increase in DA release from the SNc neuron axon varicosities/terminals in the striatum and consequent change in the levels of striatal DA
receptors, we measured striatal extracellular fluid DA levels, using
in vivo microdialysis. The results from three preliminary experiments were inconsistent (data not shown), but because the projection from the SNc to the striatum is organized in a topographic manner (Fallon and Moore, 1978), the variable results could be attributable to relatively small differences among the rats in the
location of the SNc cells subjected to gene transfer, coupled with
variations in the location of the microdialysis probe in the striatum.
Moreover, the changes in DA receptor levels that were observed (see
below) are modest in size and localized to specific areas of the
striatum, suggesting that it would be challenging to detect any
corresponding changes in DA levels; consequently, the in
vivo dialysis experiments were not pursued.
The levels of striatal DA receptors initially were assayed in
homogenates of total striatum by using receptor-binding assays. Although no changes in either striatal D1 or D2
DA receptor levels were apparent at 2 weeks after the injection of
pTHpkc into the midbrain (data not shown), analysis of the entire
striatum may have obscured regionally restricted changes in the levels
of DA receptors. Consequently, analysis via DA receptor autoradiography was undertaken, and 2 weeks after pTHpkc, pTHpkcGG, or PBS was injected into the midbrain, the rats were killed. Cresyl violet staining revealed that the injection sites were located in the posterior SNc (data not shown), which projects to the posterior striatum (Fallon and Moore, 1978). An increase in D2 DA
receptor densities, restricted to the posterior region of the striatum on the injected side of the brain, was observed in the pTHpkc group
(Table 5; this assay measures the
D2 subfamily of receptors, D2,
D3, and D4). The posterior
striatal D2 receptor binding in the pTHpkc group, but
not the control groups, differed between control and injected
hemispheres by ANOVA (p < 0.05). Furthermore, the increased D2 receptor binding in the posterior striatum
of the rats that received pTHpkc was statistically significant
(p < 0.025) in the ventrolateral and
ventromedial quadrants, but not in the dorsolateral and dorsomedial
quadrants. The increase in D2 receptor levels was modest in
size and was restricted to 2 of 12 striatal areas; consequently, if
this same increase was averaged over the entire striatum, it is
unlikely that it would be detected above background. Thus, the
apparently contradictory results from the receptor binding and receptor
autoradiography assays can be reconciled. Moreover, this localized
change in D2 receptors is consistent with both the gene
transfer that was achieved (see Stereotactic Injections and Behavioral
Testing above) and the histological results (see Fig. 3); it would have
been surprising if gene transfer to a small region of the SNc caused
changes in DA receptor levels throughout the striatum. D1
DA receptor binding was not changed in any striatal region by injection
of pTHpkc (data not shown; this assay measures the
D1 subfamily of receptors, D1 and
D5).
View this table:
[in this window]
[in a new window]
|
Table 5.
D2 DA receptor densities (fmol/mg protein) in the posterior
striatum after injection into the midbrain of pTHpkc, pTHpkcGG,
or PBS
|
|
Side effects of this HSV-1 vector system
The packaging system (Geller et al., 1990; Lim et al., 1996) used
for the behavioral experiments, KOS strain d120 virus and E5 cells
(DeLuca et al., 1985), has a low reversion frequency (<10 7) to wild-type HSV-1. After gene transfer,
all of the rats appeared healthy, exhibited unaltered ingestive
behavior and gained weight, exhibited normal motor behavior (except
when tested with apomorphine), and survived until killed. On
histological analysis, no brain tumors were observed, and there was no
evidence of reactivation of HSV-1 because no HSV-1 particle-IR was
detected at either 4 d or 4-6 weeks after gene transfer. A small
lesion area was observed around the injection site (average of 1.2 mm2 in sections containing the injection site from
four rats that received pTHlac). Unilateral destruction of SNc neurons
causes both apomorphine-induced contralateral rotational behavior and increases in D2 DA receptor levels in the ipsilateral
striatum (Ungerstedt and Arbuthnott, 1970; Creese and Snyder, 1977);
however, no such changes were observed in the vector system control
groups (pTHlac and pTHpkcGG).
| |
DISCUSSION |
A three-part genetic intervention strategy (Geller et al., 1991)
was used to modify selectively the physiology of SNc neurons with
consequent changes in apomorphine-induced rotational behavior. Our
results demonstrate that this strategy has a number of attractive features and some limitations. Use of a dominant mutation allows this
strategy to be used in wild-type animals, long-term expression supports
analysis of long-term behavioral changes, and gene transfer into adult
animals avoids potential complications from altered brain development.
The selective manipulation of a specific type of neuron can be achieved
by localized gene transfer and enhanced by using a vector that contains
a cell type-specific promoter. Post hoc histological
analysis allows for the affected neurons to be identified (although
improved detection methods are desirable). Thus, elicited behaviors can
be attributed to the altered function of identified neurons. Moreover,
although the brain often has multiple circuits that can subserve a
similar function, the selective activation of a particular
pathway in a specific group of neurons, and consequently a particular
circuit, enables analysis of the function of that particular circuit,
whereas strategies based on disrupting a specific circuit must confront
the possibility of other pathways subserving the control of that
function.
In these experiments, HSV-1 vectors were microinjected into the
midbrain, the TH promoter targeted expression to SNc neurons (Song et
al., 1997), and a constitutively active PKC produced changes in the
physiology of SNc neurons that were revealed by a DA receptor agonist.
The resulting rotational behavior was evident by 1 week and persisted
for at least 5 weeks, and the rotations were in the ipsilateral
direction. By contrast, apomorphine-induced rotational behavior was not
observed in rats that received PBS or pTHlac or even pTHpkcGG, which
encodes a single amino acid mutant of Pkc lacking PKC activity.
Constitutively active PKCs, analogous to Pkc, occur naturally.
Calpain, which is activated by the high levels of
Ca2+ produced by repetitive synaptic activation,
releases catalytic domains of PKC from the  , I, II, and isoforms (Kishimoto et al., 1989). Also, during LTP, hippocampal
neurons contain a catalytic domain of PKC that is similar to the
catalytic domains produced by calpain (Sacktor et al., 1993). Pkc
and purified rat brain PKC display similar substrate specificities, and
the catalytic domains of all PKC isoforms are highly homologous and display similar substrate specificities (Tanaka and Nishizuka, 1994).
Thus, Pkc has a structure and activity similar to those of naturally
occurring catalytic domains of PKCs.
In SNc cells, Pkc was detected in the cell body and proximal
processes, but not in the ascending axons or processes within the
striatum; in cultured neurons, Pkc usually was detected only in the
cell body and proximal processes. Different PKC isoforms are localized
to specific subcellular compartments within neurons or even specific
types of neurons, and the subcellular localization signals are thought
to be present in the regulatory domains (Tanaka and Nishizuka, 1994).
Although a similar pattern of immunoreactivity could result from the
inadequate sensitivity of the flag-IR assay, Pkc lacks a regulatory
domain, and passive diffusion seems likely to account for any transport
of Pkc from its presumed site of synthesis in the cell body, which
also would explain the observed localization of Pkc. This apparent
localization may be important for understanding how Pkc caused the
observed changes in neuronal physiology.
Pkc enhanced the activation-dependent release of catecholamines from
cultured neurons. This effect could be mediated directly by
phosphorylation of proteins that are proximal/integral to the release
machinery or indirectly by changes in the activity of distal proteins
phosphorylated by Pkc. Consistent with a direct pathway, PKC
phosphorylates specific proteins in the axon terminal, including GAP-43
(Akers and Routtenberg, 1985), and phorbol esters can increase release
from neurons or synaptosomes (Pozzan et al., 1984; Nichols et al.,
1987; Coffey et al., 1993). However, the apparent localization of
Pkc to cell bodies and proximal processes is more consistent with an
indirect pathway. There are numerous possible mechanisms for an
indirect pathway; for example, because PKC phosphorylates specific
voltage-gated ion channels, including the voltage-gated sodium channel
(Numann et al., 1991), phosphorylation of a specific voltage-gated ion
channel might alter neuronal conductivity and thereby alter release.
Although Pkc also may mediate additional changes in neuronal
physiology, the changes in neurotransmitter release that it produces
bear striking similarity to those produced by phorbol esters and by
repetitive synaptic activity (which produce PKC catalytic domains).
Also, whether direct, indirect, or a combination of mechanisms is
involved, this effect of Pkc on release is distinct from that
produced by activation of the PKA pathway: activation of the PKA
pathway by a constitutively active adenylate cyclase produced a
long-lasting increase in basal efflux without affecting K+-stimulated release (Geller et al., 1993).
Histological and pharmacological data addressed both the type of neuron
and the specific neurotransmitter system involved in mediating the
apomorphine-induced rotational behavior produced by pTHpkc. After
the injection of pTHpkc, pTHpkcGG, or pTHlac, SNc neurons were
the only midbrain catecholaminergic neurons that contained recombinant
gene products. Although some expression was observed in inappropriate
cell types in the midbrain, no expression was detected in striatal
cells. Moreover, the amount of rotational behavior correlated with the
number of SNc cells that contained Pkc. The affected
neurotransmitter system appears to have been dopaminergic because
rotational behavior was elicited by a DA receptor agonist, and this
effect was blocked by a DA receptor antagonist. Thus, we conclude that
altered dopaminergic neurotransmission from SNc neurons mediated the
observed changes in rotational behavior.
An obvious candidate for the identity and locus of altered dopaminergic
neurotransmission is an activation-dependent increase in DA release
from the striatal projections of affected SNc neurons. Pkc caused a
long-lasting and activation-dependent increase in release from cultured
sympathetic neurons, and D2 DA receptor density was
increased selectively in those striatal regions innervated by the
affected SNc neurons. This increase in DA receptor levels is consistent
with the changes that some investigators report after the
administration of drugs that increase DA release (Klawans et al., 1979;
Wilner et al., 1980), but it is not predicted by the effects of
unilateral lesion of the SNc or chronic blockade of DA receptors
(Creese and Snyder, 1977; Muller and Seeman, 1977). However,
predictions obtained from lesion experiments are not necessarily
applicable to a genetic intervention paradigm in normal rats, both
because removal of the SNc neurons causes a "gap" in the circuitry
and because other substances besides DA that also are secreted (e.g.,
peptide neurotransmitters, growth factors) are no longer present.
Furthermore, there is considerable evidence to suggest that factors in
addition to DA receptor density contribute to the direction of rotation
(Marshall et al., 1997). Moreover, Pkc may have multiple effects on
the physiology of SNc neurons that interact to influence the direction
of rotation. For example, SNc neurons can release DA from their cell
soma and dendrites, and it is possible that such additional sites also
may be involved.
The presence of Pkc in a small percentage (~0.1-2%) of SNc
neurons was sufficient to alter apomorphine-induced rotational behavior. Consistent with this observation, in normal animals the small
imbalances in the nigrostriatal system between the two hemispheres
appear to underlie spontaneous rotational behavior (Jerussi and Glick,
1976; Glick et al., 1981). Also, because the flag-IR positive cells
were in the posterior SNc, the local concentration of affected neurons
was probably higher than the overall average. Such localized
concentrations of affected SNc neurons may be critical for causing both
the localized changes in striatal D2 DA receptor levels and
the change in apomorphine-induced rotational behavior.
pHSVpkc did not alter apomorphine-induced rotational behavior, and
this may be because the majority of the expression (~95%, similar to
results with pHSVlac) was in glia and in non-SNc neurons. Because glia
do not take up, store, or release catecholamines (Hansson, 1983),
activation of PKC pathways in glia seems an unlikely direct effector of
changes in dopaminergic neurotransmission. However, Pkc in the
non-SNc neurons could alter their physiology to counterbalance the
effects of Pkc in the SNc neurons on apomorphine-induced rotational
behavior. For example, pHSVlac and pHSVpkc supported significant
expression in SN reticulata (SNr) cells with neuronal morphology.
Because SNr neurons that project to the SNc are GABAergic, it is
conceivable that Pkc enhances GABA release at these synapses, thereby counterbalancing the effects of Pkc in SNc neurons.
Our results demonstrate that the three-part genetic intervention
strategy used here has promise, but a number of questions remain
unanswered. This strategy is critically dependent on the vector system,
and the HSV-1 vector system used in this study is not without its
problems, especially cytopathic effects and immune responses, caused
primarily by the helper virus (Wood et al., 1994). A recently developed
helper-virus-free packaging system has addressed many of these concerns
(Fraefel et al., 1996); however, additional improvements are still
required. Although it remains to be seen if this strategy can be
applied to more complex behaviors, such as learning and memory, each of
the established strategies for modifying brain physiology (including
transgenic mice, cell transplantation, and surgical, pharmacological,
and electrophysiological interventions) also has particular strengths
and weaknesses. As such, the present strategy may prove to be equally
useful.
| |
FOOTNOTES |
Received March 23, 1998; accepted March 25, 1998.
This work was supported by the Pharmaceutical Manufacturers Foundation
(S.S.); Council for Tobacco Research USA Grant 3129 (H.R.); National
Institutes of Health (NIH) Grant NS25143 and Research Scientist
Development Award MH00967 (J.H.); NIH Grant HD24236 (R.N.); NIH Grant
AG10827 (A.G. and K.O.); NIH Grant NS34024, Neurovir, National Alliance
for Research on Schizophrenia and Depression, and the Burroughs
Wellcome Fund (A.G.). We thank Drs. N. DeLuca for both d120 virus and
E5 cells, J. Knopf for the rat PKCII cDNA, M. Leahy for the mouse
anti-flag antibody, M. D. Browning for both the purified rat brain
PKC and the constitutively active fragment obtained by partial
digestion with trypsin, and A. Heller for MN9D cells. We also thank Ms.
Courtney S. Holmes for technical assistance. We gratefully acknowledge
Dr. Dean M. Hartley for constructing the first pTHpkc vector and for
helpful discussions with Dr. Song.
This manuscript is dedicated to the memory of Dr. Francis O. Schmitt
(1903-1995).
Correspondence should be addressed to Dr. Alfred Geller, Division of
Endocrinology, Children's Hospital, Boston, MA 02115.
| |
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Abstract
Introduction
