Division of Biology, California Institute of Technology, Pasadena, California 91125
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ABSTRACT |
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Nadeau, H.,
S. McKinney,
D. J. Anderson, and
H. A. Lester.
ROMK1 (Kir1.1) Causes Apoptosis and Chronic Silencing of
Hippocampal Neurons.
J. Neurophysiol. 84: 1062-1075, 2000.
Lentiviral vectors were constructed to express the
weakly rectifying kidney K+ channel ROMK1 (Kir1.1), either
fused to enhanced green fluorescent protein (EGFP) or as a bicistronic
message (ROMK1-CITE-EGFP). The channel was stably expressed in cultured
rat hippocampal neurons. Infected cells were maintained for 2-4 wk
without decrease in expression level or evidence of viral toxicity,
although 15.4 mM external KCl was required to prevent apoptosis of
neurons expressing functional ROMK1. No other trophic agents tested
could prevent cell death, which was probably caused by K+
loss. This cell death did not occur in glia, which were able to support
ROMK1 expression indefinitely. Functional ROMK1, quantified as the
nonnative inward current at 144 mV in 5.4 mM external K+
blockable by 500 µM Ba2+, ranged from 1 to 40 pA/pF.
Infected neurons exhibited a Ba2+-induced depolarization of
7 ± 2 mV relative to matched EGFP-infected controls, as well as a
30% decrease in input resistance and a shift in action potential
threshold of 2.6 ± 0.5 mV. This led to a shift in the relation
between injected current and firing frequency, without changes in spike
shape, size, or timing. This shift, which quantifies silencing as a
function of ROMK1 expression, was predicted from Hodgkin-Huxley models.
No cellular compensatory mechanisms in response to expression of ROMK1
were identified, making ROMK1 potentially useful for transgenic studies
of silencing and neurodegeneration, although its lethality in normal
K+ has implications for the use of K+ channels
in gene therapy.
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INTRODUCTION |
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Potassium ion selective channels establish the neuronal resting membrane potential (RMP) and restore it after firing. We describe a method for expressing a new set of K+ channels in neurons, to alter their excitability. Neuronal RMP is often 10-20 mV depolarized relative to the K+ reversal potential, and thus more open K+ channels are expected to hyperpolarize the cell. Moreover, more open channels lead to a reduction in input resistance, rendering excitatory synaptic currents less effective in depolarizing the cell to threshold. In brief, a change in input resistance due to more open K+ channels can abolish action potential firing ("silence" the cell).
There are several reasons to silence neurons in vitro and in vivo.
Selectively targeting pathways or populations of neurons has revealed
the importance of interneuronal communication in developing
(Murakami et al. 1992) and adult (de la Cruz et
al. 1996
) systems. Botulinum toxin is used to lessen muscle
spasms in patients with cerebral palsy (Flett et al.
1999
) and spinal cord injuries (Al-Khodairy et al.
1998
), and targeted lesions are often the only possible
treatment for those with intractable epilepsy (Jallon
1997
; Nayel et al. 1991
). Control of
excitability may also be important for lessening neurological damage
following ischemic injury or in degenerative disease (Rodriguez
et al. 1998
). A genetic approach has two major advantages over
toxins and surgery: one, it can target a pathway that is not fully
understood; and two, it has the potential to be fully inducible and
reversible over a time course of hours.
However, to evaluate genes as silencing candidates, it is important to be able to translate alteration of excitability seen in culture into in vivo long-term behavior. We use an HIV-based lentiviral vector to create an in vitro model of transgenesis, with a K+ channel as the candidate silencer. Neurons are transduced soon after plating and allowed to grow and develop for days to weeks with unopposed channel expression. The viral genome is integrated into its host at low copy number, leading to stable expression levels and no interference with host protein synthesis. Efficacy and lack of toxicity of the viral vector itself are established, and no inactivation or down-regulation of the channel is observed over a 3-wk period. However, the chronic efflux of K+ leads to apoptotic cell death unless counteracted by a raised K+ concentration in the culture medium. This imposes limits on the possible in vivo use of ROMK1, and perhaps of other K+ channels.
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METHODS |
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Molecular biology
The Xba I fragment of pEGFP (Clontech, Palo Alto, CA), containing the complete enhanced green fluorescent protein gene, was inserted into pTRE (Clontech) in the correct orientation to give pTRE-EGFP. This was cut with EcoR I and Age I, and a 33-bp polylinker containing Pac I and Swa I sites
5'AATT CCCCC TTAATTAA CTAG ATTTAAAT CCCA
3' GGGGG AATTAATT GATC TAAATTTA GGGTGGCC
was linked to the sticky ends. The cap-independent translation enhancer (CITE) (500 bp from encephalomyocarditis virus) was amplified by polymerase chain reaction (PCR) (High Fidelity PCR Kit, Boehringer Mannheim, Indianapolis, IN) from pCITE-2a (Novagen, Madison, WI) and inserted in frame into the Age I-Nco I sites. PCR was verified by complete sequencing on both strands. The fragment including the polylinker and CITE-EGFP was then subcloned into the EcoR I-Not I sites of pCITE-2a to give [pCITE-(CITE-EGFP)].
The ROMK1 complete cDNA (Ho et al. 1993) (1.3 kb) plus
0.9 kb of 5' untranslated sequence was excised from pSport (Life
Technologies, Gaithersburg, MD) as a single Mlu I fragment
and inserted into the BssH II site of pNEB193 (New England
Biolabs, Beverly, MA) to give pNEB193-ROMK1; the orientation in which
the Pac I site was located on the 3' end of the gene was selected.
To create lentiviral constructs, sequences were cloned into the plasmid pHR' (gift of Didier Trono, The Salk Institute), which contains a human cytomegalovirus (CMV) promoter. The 1.3-kb CITE-EGFP was removed from pCITE-(CITE-EGFP) with BamH I and Xho I and ligated to the corresponding sites of pHR' to give pHR'CITE-EGFP. The EcoR I and Pac I sites on the 5' end of this fragment serve as a unique polylinker in this vector; ROMK1 was inserted into the EcoR I and Pac I sites after removal from pNEB193-ROMK1 to give pHR'ROMK1-CITE-EGFP (Fig. 1A).
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The control construct, encoding for EGFP alone, was constructed by subcloning the Xba I fragment of pEGFP into pBluescript (Stratagene, La Jolla, CA). The gene was then excised with BamH I and Xho I and ligated into the same sites of pHR'.
To generate the EGFP-ROMK1 fusion protein, the EGFP gene minus the final stop codon was amplified from pEGFP by PCR, generating BamH I and Mlu I ends; this was inserted into the corresponding sites of pHR' to give pHR'EGFPnostop. The first 500 bp of ROMK1 was also amplified by PCR, giving an Mlu I-Bgl II fragment. The remaining 1.7 kb was excised from pSport with Bgl II and Xho I, and a three-way ligation between these two fragments and pHR'EGFPnostop (cleaved with Mlu I and Xho I) produced the final construct, pHR'EGFP-ROMK1 (Fig. 1B). All PCR products were verified by complete sequencing, and ligations were verified by restriction digest and partial sequencing.
Generation of lentiviruses
Plasmids were amplified using Maxi and Mega kits from
Qiagen (Valencia, CA). The plasmids pHR'ROMKI-CITE-EGFP,
pHR'EGFP-ROMK1, pHR'EGFP, or pHR' alone (which encodes LacZ) were
cotransfected into 293T cells with the plasmids pMD.G and
pR8.9 in the ratios published (Naldini et al. 1996
).
Transfection was performed in 15-cm tissue culture dishes (Falcon,
Oxnard, CA) at 80-90% confluence, using the cationic lipophilic
reagents Superfect or Effectene (Qiagen). A total of 20 µg of plasmid
DNA was added per 15-cm dish when Superfect was used; a total of 4 or 8 µg was added with Effectene. In the latter case, the ratio of
Enhancer to microgram of plasmid DNA was always maintained at 8:1.
Supernatant was harvested at 48, 72, and 84 h after transfection
and spun at 1,000 g for 5 min to remove cellular debris. It
was then passed through a 0.45 micron filter and either subjected to
two 15-min spins at 1,000 g in a Centricon-500 unit
(Millipore, Bedford, MA) (ultrafiltered virus) or spun for 1.5 h
at 4°C in a Beckman ultracentrifuge, using an SW41.1 Ti
swinging-bucket rotor at 20,000 rpm (ultraconcentrated virus). Virus
from either stock was assayed for infectivity on 293 T or Chinese
hamster ovary cells. Ultrafiltered virus averaged 5 × 106 transducing units (TU)/ml, ultraconcentrated,
5 × 107. The former was stored in 1 ml
aliquots and the latter in 50 µl aliquots, all at
80°C.
Cell culture
Pregnant Wistar rats were euthanized by inhalation of CO2 at day 18 of gestation. Embryos were immediately removed by caesarian section, and hippocampi rapidly extracted under stereomicroscopic observation under sterile conditions, cut into 1 mm pieces, and digested with 0.25% trypsin and 0.25 mg/ml DNAse (Sigma, St, Louis, MO) at 36°C for 15 min. The pieces were then gently rinsed in Hanks' balanced salt solution without Ca2+ or Mg2+ (HBSS, Life Technologies), washed twice in plating medium, and gently triturated in 1 ml of plating medium with five passes through the 0.78 mm opening of a tip of a P-1000 Pipetman. Suspended cells were removed with a Pasteur pipette, and the remaining pieces triturated once more. The resulting suspensions were gravity-filtered through a 70-µm nylon mesh to remove large debris, and centrifuged for 2 min at 150 g to pellet the cells, which were resuspended by trituration as above. Approximately 35,000 cells were plated in an area 15 mm in diameter at the middle of a 35-mm plastic culture dish that had been coated with poly-D-lysine (PDL) and laminin. Cultures were maintained at 36°C in a 5% CO2 incubator. One half volume of medium was changed twice weekly with culture medium. Plating and feeding medium was Neurobasal with B27 supplement, with 500 µM Glutamax, 25 µM glutamate, and 5% horse serum (Life Technologies); [K+] in this medium is 5.4 mM.
HEK 293 cells were maintained with weekly passages in DME high glucose medium (GIBCO) supplemented with 10% fetal bovine serum, 200 µM glutamine, and penicillin/streptomycin. Transfections were performed with Effectene (Qiagen) in 35-mm dishes according to the manufacturer's instructions.
Infection of hippocampal neurons
Cells were infected 1-3 d after plating by the addition of 20-30 µl of ultraconcentrated or 200-300 µl of ultrafiltered virus to the medium. Viral supernatant was not washed off, although cells continued to be fed on a weekly basis. Stocks of virus were pooled so that each dish in a preparation received the same concentration. For "high K+" cells, supplementary KCl was added to a total concentration of 15.4 mM 12-24 h after infection. At least 40 h were allowed to elapse before assaying for EGFP expression. Control neurons were matched for age (to within 1 d) and for time since application of drugs and/or EGFP-only virus. All controls were from separate dishes; nonfluorescent cells from ROMK1 dishes were never used as controls because of the difficulty of excluding faint fluorescence. Sixteen of thirty-two high K+ controls were EGFP-infected and 16/32 were uninfected; data from these two groups were pooled when no effects of infection were detected on resting membrane potential, spike threshold, input resistance, or response to Ba2+. Similarly, 3/9 low K+ controls were EGFP-infected, 6/9 uninfected, and the data pooled. Data from cells in high versus low K+ were never pooled. When pooled data were averaged, the control and ROMK1 groups contained equal proportions of neurons with identical ages postinfection and postplating. All data presented for ROMK1-infected neurons are from those infected with the bicistronic message; the fusion protein was used only in localization studies and HEK cell recordings, primarily because its fluorescent signal was fainter than that of the cytosolic EGFP. EGFP was visualized for electrophysiology by fluorescence microscopy on a Nikon inverted microscope illuminated by a 100 W Hg lamp, using either a EndowGFP double band-pass filter (exciter 470/40; dichroic 495 LP; emitter 525/50) or a HiQFITC band-pass/longpass filter (exciter 480/40; dichroic 505 LP; emitter 535/50) (Chroma Technologies, Brattleboro, VT). The objective was a 40× air with NA = 0.5 (Nikon, modified by Modulation Optics Inc., Greenvale, NY). Confocal images of fixed specimens stored in phosphate-buffered saline (PBS) were obtained on a Zeiss LSM-510 using a laser tuned to 488 nM for FITC and EGFP and a 40× Plan-Neofluar water immersion lens with NA = 0.9 (Zeiss). Beta-galactosidase was visualized by standard methods after fixation for 2 min in 50/50 methanol/acetone. No viral toxicity was observed with either control construct (EGFP or LacZ).
Whole-cell recording
All recordings were performed at room temperature. For
hippocampal neurons, borosilicate dot glass capillaries (Sutter
Instruments, Novato, CA) were pulled to a tip resistance of 5-10 M
and filled with either a Mg2+-containing internal
solution consisting of (in mM) 100 K-gluconate, 10 HEPES, 3 phosphocreatine, 1.1 EGTA, 3 MgATP, 0.2 NaGTP, 5 MgCl2, 0.1 CaCl2; or a
Mg2+-free internal solution, 100 K-gluconate, 10 HEPES, 1.1 EGTA, 0.1 CaCl2, both adjusted to pH
7.2 with KOH and 250 mOsm with sucrose. The bath solution contained (in
mM) 110 NaCl, 10 HEPES, 5.4 KCl, 1.8 CaCl2, 0.8 MgCl2, 10 glucose, adjusted to pH 7.4 with NaOH
(EK =
76 mV at 25°C). Calculated
junction potential for these solutions is 14 mV (Clampex 8.0), and all
reported membrane potentials are corrected for this value. Tetrodotoxin
(TTX) was bath-applied to a final concentration of 1 µM or perfused
at 500 nM; Ba2+ (500 µM),
Co2+ (1 mM), and a "hippocampal cocktail"
consisting of bicuculline (10 µM), APV (50 µM), and CNQX (20 µM)
(all from RBI, Natick, MA) were perfused continually through flow pipes
of 250 µm internal diameter mounted ~500 µm from the recorded
cell. The dish was washed with 4-6 ml of control saline between
recordings. For HEK 293 cells, capillaries were pulled to a tip
resistance of 2-5 M
and filled with an internal solution containing
(in mM) 130 KCl, 0.8 MgCl2, 5 EGTA, 5 MgATP, 10 HEPES (pH to 7.2 with KOH); the bath solution consisted of (in mM) 137 NaCl, 10 HEPES, 4.0 KCl, 1.8 CaCl2, 1.0 MgCl2, 10 glucose, adjusted to pH 7.4 with NaOH
(EK =
91 mV at 25°C). Junction
potential with this solution is 4.5 mV.
Signals were recorded with an Axopatch 1D amplifier (Axon Instruments,
Foster City, CA) and sampled by a Digidata 1200 at 20 kHz (100 kHz for
transients) to a Pentium PC. Series resistance was not
compensated but was monitored throughout the recording. For
I-V analysis, series resistance was compensated off-line
(Nadeau and Lester 2000; Traynelis 1998
).
Current and voltage commands and data acquisition were performed using
PCLAMP6.0 and 8.0. Data were analyzed with AxoGraph 3.5 (Axon);
individual compiled modules were written using CodeWarrior Pro
(Metrowerks Software). Cells were eliminated from the analysis if
series resistance changed by more than 20% during the course of the
recording, if the cell spontaneously depolarized, or if the membrane
capacitance changed in either direction by more than 10%. Data from
cells in TTX and cocktail were pooled for many analyses as there were
no detectable differences in membrane potential or effects of ROMK1
expression or blockade. Data on threshold and silencing were
unavailable for cells with TTX bath application (n = 10 ROMK1 cells, 5 high K+ controls).
Immunocytochemistry
TUNEL staining was performed with the In Situ Cell Death Detection Kit-POD (Boehringer) according to the manufacturer's instructions; the horseradish peroxidase (HRP)-conjugated secondary was developed with nickel-diaminobenzidine reagent and visualized under brightfield. Antibody staining was with polyclonal rabbit anti-ROMK1 (Alomone Labs, Jerusalem, Israel). Cultured cells were fixed for 2 min in methanol/acetone, permeabilized for 2 min on ice with 1% Triton X-100, and preincubated for 30 min in PBS with 5% goat serum. Primary antibody was diluted 1:50 in the same solution and incubated with gentle shaking for 2 h at room temperature or overnight at 4°C. The dish was washed five times and incubated with fluorescent secondary antibody (Cy-3 conjugated goat anti-rabbit, Jackson ImmunoResearch, West Grove, PA) for 60 min at 37°C and visualized with a Texas Red filter (exciter 560/55; dichroic 595 LP; emitter 645/75) (Chroma). Omission of primary antibody led to weak, nonspecific staining. Fluorescence was quantified with NIH Image using confocal images to distinguish membrane-bound from cytoplasmic fluorescence.
Pharmacology
Drugs were added to infected neurons 12-24 h after application
of virus. All were dissolved in stock solutions in DMSO in the
following concentrations (in mM): 10 BayK 8644 (Calbiochem, La Jolla,
CA); 10 nifedipine (ICN Biomedicals, Aurora, OH); 2 thapsigargin (RBI,
Natick, MA); 100 8-4(chlorophenylthio)cAMP (cpt-cAMP, Boehringer).
Brain-derived neurotrophic factor (BDNF) (Sigma) was dissolved in PBS
containing 0.1% bovine serum albumin to 10 µg/ml and stored in
aliquots at 20°C.
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RESULTS |
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Transient transfection of bicistronic and fusion vectors
Transfected into HEK 293 cells, the vectors pHR'ROMK1-CITE-EGFP and pHR'EGFP-ROMK1 produce weakly inwardly rectifying currents that are blockable by 500 µM Ba2+ in a time- and voltage-dependent manner. The block is reversible on Ba2+ wash-out and reverses at EK after leak subtraction (Fig. 2). The bicistronic message produces a strong fluorescent signal throughout the cell, while the fusion protein appears as a weaker, punctate fluorescence mostly directed to the plasma membrane and completely excluding the nucleus (data not shown; see Fig. 6 for distribution in neurons).
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Effects ROMK1 on neuronal morphology and survival
Hippocampal neurons infected with control virus, bearing EGFP only, become visibly fluorescent after 24-36 h and increasingly so for several days thereafter. Cell morphology, processes, and underlying glia are unchanged. Infected cells can be maintained and recorded from for up to 6 wk; their electrophysiological properties are identical to those of normal cells (Fig. 3, A and B; Table 2). Hence, there is no viral toxicity detected within the sensitivity of our experiments.
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In contrast, cells infected with ROMK1-CITE-EGFP show no healthy green
neurons at 48 h. Expression is limited to astrocytes, possibly
activated microglia, and dead or dying cells of varying morphologies.
Many dead cells are shrunken and floating, a classic indicator of
apoptosis (Gibson 1999) (Fig. 3, C and
D). TUNEL staining (Villalba et al. 1997
)
reveals no apoptotic cells 24 h post infection, but nearly 100%
of the neurons are apoptotic at 48-72 h (Fig. 3,
E-G). By the fourth to fifth day, few neurons remain in the dish (Fig. 3H). Glial cells can maintain
stable infections with ROMK1-CITE-EGFP for weeks or months, as can CHO or HEK-293 cells. The latter may be serially passaged indefinitely without visible change in the proportion of fluorescent cells. Thus
cultured hippocampal neurons but neither glia nor clonal cell lines
appear susceptible to overexpression of ROMK1.
Channel block by inorganic ions is ineffective at rescuing
ROMK1-expressing neurons. Dishes supplemented with 200-500 µM
BaCl2 show the same pattern of apoptosis as
untreated cultures; however, the blockade at these concentrations
affects mainly inward and not outward K+ currents
(Ho et al. 1993). Higher concentrations of
Ba2+ permit <10% of ROMK1-expressing neurons to
survive but are toxic to glia and hence lead to massive deterioration
in all cells.
Elevated K+ prevents apoptosis
We tested the hypothesis that K+ loss
through ROMK1 causes apoptosis. When the neuronal growth medium
K+ concentration is increased to 15.4 from the
usual 5.4 mM (high K+), which shifts
EK from 79 to
51 mV at 36°C,
fluorescence microscopy of ROMK1-CITE-EGFP infected cells at 48-72 h
reveals 50-90% green neurons with normal morphology (Fig. 3,
I and J). Smaller elevations of
K+ levels, to 9.4, 11.4, and 13.4 mM, are
insufficient to maintain healthy infected cells (data not shown).
Importantly, increased K+ has no visible effect
on the fluorescence of EGFP-only cells. ROMK1-expressing neurons can be
maintained with weekly feedings of medium containing 15.4 mM
K+ for >3 wk. This life span is similar to that
of controls in high K+. Thus this elevation of
K+ is sufficient to prevent apoptosis and to
restore ROMK1-infected neurons to an apparently normal state of health.
The results are consistent with a specific action of
K+: activation of apoptotic pathways due to
K+ efflux (Yu et al. 1997).
However, they may be equally explained by nonspecific trophic actions
of chronic depolarization and Ca2+ entry. The use
of Ca2+ agonists and antagonists is necessary to
distinguish these mechanisms.
The protective effect is specific to K+
Unlike many neuronal cells in culture, hippocampal neurons
normally are not dependent on sustained depolarization for survival. They may be grown in media containing as little as 2.5 mM
K+ or in 1 µM TTX, a concentration sufficient
to block all action potentials (Table 1).
On the other hand, they are highly sensitive to excitotoxicity, and
increased current through L-type Ca2+ channels is
harmful (Porter et al. 1997). It is therefore unlikely that any amount of ROMK1-mediated hyperpolarization and altered membrane conductance could lead to loss of Ca2+
sufficient to cause the rapid, total cell death that we observe. To
eliminate this possibility, however, we treated infected dishes with
combinations of trophic factors and Ca2+ agents:
cpt-cAMP (0.1-0.5 mM); thapsigargin (100 nM), which causes the release
of intracellular Ca2+ stores; BayK 8644 (1 µM),
an L-type Ca2+ channel agonist; BayK 8644 plus
nifedipine (10-100 µM), an L-type antagonist expected to counteract
the effects of BayK; glutamate (40 µM); or BDNF (20 ng/ml).
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None of these agents are able to maintain ROMK1-infected neurons (Table 1). Neurons in normal K+ exposed to agents that increase intracellular Ca2+ show 100% apoptosis by 48 h, as in untreated cells (Fig. 4A). Neither EGFP-infected controls nor ROMK1 cells in high K+ are harmed by these drugs. Ca2+ antagonists reverse the beneficial effects of high K+ only slightly.
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There is a difference in appearance between the ROMK1-infected cells
with and without Ca2+ elevation. In the former
case, nearly every neuron in the dish is fluorescent, but their
morphologies are highly bizarre; they are completely depolarized, and
they detach from the dish (Fig. 4B). In the latter case, the
neurons disappear before many distorted forms are seen. This suggests
that Ca2+ elevation is acting to prevent
phagocytosis by astrocytes or microglia, without reversing the lethal
phenotype caused by the channel; this could result either from effects
of elevated Ca2+ on the apoptotic process
(Bratton et al. 1999) or from direct effects on the glia.
These data point to K+-efflux mediated apoptosis as the cause of cell death in long-term ROMK1 expression. Loss of K+ may impose a metabolic burden on the cell, instead of or in addition to triggering apoptotic pathways; this is a topic for future study.
K+ loss is a critical feature of ROMK1 expression that restricts its applicability for gene therapy or transgenics. However, it may still possess properties of a useful silencing gene, and understanding its effects can lead to improvements in silencing strategies. In addition, outward ROMK1 currents may not occur in all neurons at all ages, especially those subjected to chronic electrical input and depolarization. It is therefore informative to examine the properties of cells maintained in high K+, where ROMK1-infected cells have the same growth patterns, morphology, and life span as matched EGFP-infected controls.
Electrophysiology: Expression levels and localization
Because this study concerns the effects of a new conductance on
encoding, an accurate description of membrane parameters is important.
Many neurons in culture can be successfully described by a single
capacitance, input resistance, and series (pipette) resistance. The
resistance of the proximal dendrites is large enough that little
attenuation occurs in the signal as it travels to the soma, and the
soma resistance is itself large enough to overcome most of the
artifacts caused by the series resistance (typical values of input and
series resistance are 1000 and 10 M, respectively). However, for
cells infected with a K+ channel, this may no
longer be the case: input resistance may decrease two- to fivefold,
with a corresponding increase in artifacts. We therefore use a
two-compartment model to provide better exponential fits and the
ability to distinguish somatic conductances from poorly space-clamped
conductances on the distal dendrites (APPENDIX B; see also
Nadeau and Lester 2000
).
ROMK1-infected and control cells were similar in age and size of
both the proximal and distal compartments (Table
2); the apparently small size of
CD in the low K+
controls is due to a preponderance of young cells in this group [only
2 of 9 were over 7 days in culture (dic)]. The cells are somewhat
smaller than in other studies of hippocampal neurons (Mennerick
et al. 1995), which primarily reflects our inclusion of young
cells, although some stunting due to high K+ may
also occur.
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Blockade by 500 µM Ba2+ was used to quantify
expression. The honeybee toxin tertiapin (Jin and Lu
1998) has been proposed as a specific blocker for inward
rectifier K+ channels, but it is sensitive to
oxidation and its effects on neurons are unknown. Application of fully
oxidized tertiapin at a concentration of 2 kD
(kD = 7.7 µM) led to increased leak
in 2/3 ROMK1 cells and 3/3 controls (not shown). Washout was slow and
incomplete. While a stable version of the venom has recently been
synthesized (Jin and Lu 1999
), its effects on normal
neurons will have to be investigated before it becomes a useful tool.
On the other hand, inhibition of the channel by submillimolar
Ba2+ is well characterized, rapid, and easily
reversible (Choe et al. 1998; Ho et al.
1993
). Its only drawback is incompleteness of block at
depolarized potentials, following the Woodhull equation
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Total blocked current at a holding potential of 130 mV (corresponding
to a membrane potential of
144 mV) (Fig.
5A) shows an upward trend with
time in ROMK1-infected neurons, with levels of expression peaking at
6-7 d and remaining steady thereafter (Fig. 5B). In low
K+ controls, this concentration of
Ba2+ blocks no native channels, but after several
days in high K+ up-regulation of native inwardly
rectifying K+ channels (Guo et al.
1997
; Knutson et al. 1997
) causes high
K+ controls to exhibit a smaller but significant
Ba2+ block. It is not certain that the
up-regulated channels are members of the Kir superfamily, but for
convenience, we shall abbreviate them as EIRKs, for "endogenous
inwardly rectifying K+ " channels. We show
(APPENDIX A, Fig. 8) that they display strong inward
rectification. Levels of this channel(s) remain essentially constant
over the period studied.
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With Mg2+-free internal solution, the mean
current at this voltage is 1.75-fold that is seen with internal
solution containing Mg2+, ATP, and GTP
(n = 5 for Mg2+-free cells,
P < 0.15, t-test), consistent with
suppression of ROMK1 by cytoplasmic ATP (Ho 1993). If
blockable currents in control cells are subtracted from those in ROMK1
neurons (this slightly underestimates the amount of ROMK1 expressed,
see APPENDIX A), the resulting average expression level is
approximately one quarter of that previously measured with
adenovirus-mediated G-protein-activated inward rectifier K+
channel expression (Ehrengruber et al. 1997
).
The maximum level in the present experiments equals the mean of 40 pA/pF as reported by Ehrengruber et al.; a level this high represents
an unusual situation for a retrovirus and is probably due to positional
effects (Schubeler et al. 1998
). However, even with
retroviral vectors, current densities of >12 pA/pF were not unusual,
representing 11% of all infected cells.
We identify a subgroup of infected neurons that do not fire action potentials at the maximum tested level of current injection (200 pA), but exhibit normal firing patterns after blockade by Ba2+. These are designated "completely silenced cells" and show significantly greater blockable currents than the mean of all ROMK1-infected neurons; these cells illustrate the effects of the maximal level of expression achievable with lentiviral transduction.
Localization was investigated by antibody staining, confocal imaging of EGFP-ROMK1 fusion expression, and two-compartment calculation of currents (Fig. 6). No specific localization pattern was identified, consistent with simple diffusion of proteins throughout the cell.
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Effects of ROMK1 on encoding properties: Change in RMP with Ba2+
Full I-V relations with and without
Ba2+ demonstrate voltage dependent block in both
ROMK1 and control neurons (Fig. 7). While EIRK complicates the picture at hyperpolarized potentials, blockable currents near typical RMP values of 60 to
65 mV are due almost entirely to ROMK1, giving an I-V similar to that seen in HEK
cells (Figs. 7E and 8).
Therefore, a significant change in RMP is expected in expressing cells
with the addition of Ba2+, but not in controls.
This depolarization can be predicted, with two assumptions: that
Na+ and K+ are the only
ions contributing to RMP, and that Na+
conductance does not change with Ba2+. Then
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With increasing periods, postinfection changes in RMP become
smaller, even in cells expressing 20-40 pA/pF of
Ba2+ blockable current at hyperpolarized
potentials. At 13 dpi, average Vm has
fallen to
5.1 ± 1.0 mV (n = 12), even as ROMK1
expression has increased (Fig. 5B). There are two
identifiable reasons for this. First,
VmBa is hyperpolarized by 1 mV, from
58 ± 1 to
59 ± 2 mV, making it closer to
EK; the predicted change has now
decreased slightly, to
7.0 ± 1.6 mV.
In addition, there is now a discrepancy between the predicted and
observed values. Its origin is a hyperpolarization resulting from
Ba2+ wash-in in neurons in long-term high
K+; this means that our assumption that
GNa does not change with Ba2+ is no longer correct. At 13 dpi, the mean in
controls is 1.5 ± 1 mV, n = 10. Subtracting this
from the mean given by Eq. 2 gives an adjusted
value of 5.5 ± 1.9 mV, well in line with the observations.
By 18-21 dic, average RMP change in ROMK1 cells has fallen to
2.4 ± 0.8 mV (n = 12). This can be almost
entirely explained by proximity of
VmBa to
EK: cells are now hyperpolarized to
66 ± 1 mV, and the predicted change is
3.5 ± 0.6 mV.
The discrepancy of 1.8 ± 0.8 mV results from 4/12 cells that show no depolarization or even hyperpolarization in
Ba2+ (2 cells with hyperpolarization, 1.9 and 3.2 mV).
In controls (n = 7), hyperpolarization is also
seen only in a subset of cells, always accompanied by the slow,
stepwise block shown in the last panel of Fig. 5A. Mean
change in RMP is 4.66 ± 2.2 mV, range 1.2 to 15 mV, with no
hyperpolarization seen in two cells and
10 mV in two cells. This may
reflect different subpopulations of neurons with differing responses to
elevated K+ and is a topic for future study. The
degree of hyperpolarization does not appear to correlate with level of
ROMK1 expression, but the numbers of neurons displaying this phenomenon
are too small for meaningful statistics.
These compensatory channels may explain the extended lifetime of infected cells washed back into normal K+ after >7 dpi: 4-5 d versus <48 h for cells infected in low K+. However, despite the disappearance of membrane hyperpolarization, silencing is not eliminated: the effects of the channel on excitability increase with increasing levels of ROMK1 expression and are only partially dependent on RMP.
Three changes are responsible for silencing
In the cells we studied, spiking elicited by current injection shows a pattern typical of hippocampal neurons (Fig. 9A). There is a definite threshold potential for the occurrence of spikes (threshold was taken to be the point at which the time derivative of the current exceeded 10 V/s). In most cells, accommodation of firing rates occurs during a depolarization lasting 800 ms. Higher currents increase the firing rate, but beyond a maximum level of ~20 Hz, spikes broaden and firing rates decrease. Plots of spike frequency versus current injection reveal that the addition of Ba2+ to infected cells changes the threshold of spike response without affecting the overall shape of the frequency-versus-current relation (Fig. 9B). We define this shift, with the dimensions of pA, as the measure of "silencing," S.
|
S can be broken down into three components, indicating the
amount of current necessary to overcome the ROMK1-induced
hyperpolarization Vm = Vm
VmBa, the increased membrane
conductance
G = G
GBa, and the change in threshold
T = T
TBa
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(3) |
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(4) |
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(5) |
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Since cells must fire both with and without Ba2+ to yield values of S, the completely silenced cells are excluded from this analysis. Nevertheless, even neurons expressing an average amount of ROMK1 have significant Ba2+blockable conductance at depolarized potentials, and the effects on firing are well predicted by this simple ohmic model (Fig. 11).
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DISCUSSION |
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K+ channels have been proposed as candidates
for transgenesis and gene therapy, because expression of several
different types has been shown to decrease excitability in cultured
neurons (Ehrengruber et al. 1997; Johns et al.
1999
). In all of these experiments, however, the channels were
active only in the presence of agonist or inducer. Expression was
activated during or immediately before electrophysiological recording,
so that long-term health effects of the gene or neuronal compensatory
mechanisms could not be identified. Furthermore, adenovirus expression
levels are many times those seen in transgenic animals; it is unknown
whether a small change in K+ conductance will
have equally profound effects. In fact, mice transgenic for Shaker
(AKv1.1a) (Sutherland et al. 1999
) show a hyperexcitable
phenotype, with spontaneous EEG discharges and lowered seizure
thresholds, in apparent contradiction to the above results and
underlining the complexity of the problem.
We begin to address some of these issues by examining chronic, unopposed K+ conductance expressed at low levels from a viral vector with minimal toxicity. The weak inward rectifier ROMK1 results in apoptosis in 100% of dissociated hippocampal neurons, independent of age. Cell death cannot reliably be prevented by any pharmacological agents other than K+ that we have studied. Ca2+ channel agents and growth factors can block apoptosis, but the resulting neurons are depolarized and morphologically abnormal. Accordingly, K+ rescue is not antagonized by Ca2+ channel blockers.
Although increasing K+ to 15.4 mM is sufficient
to permit survival of nearly all ROMK1-expressing cells, this
manipulation has profound effects, especially after prolonged times.
EGFP-infected and uninfected cells up-regulate native inward
rectifiers, which we have termed EIRKs in this paper, so that they are
hyperpolarized by 5 ± 2 mV when recorded in normal
K+ medium. They also show a threefold increase in
current at 144 mV and a slightly more than twofold decrease in input
resistance at
94 mV relative to neurons in normal
K+. After more than 2 wk in culture, additional
channels begin to appear, some of which (a) are permeable to
Na+ and (b) lead to hyperpolarization on
Ba2+ application (details in APPENDIX
A).
Application of 500 µM Ba2+ leads to block of EIRK and ROMK1 at hyperpolarized potentials, while at more depolarized potentials near threshold, the EIRK conductance disappears and a smaller fraction of ROMK1 is blocked. However, the blockade is sufficient to restore the membrane potential, spike threshold, and whole-cell conductance of ROMK1 expressing cells to near control values while they are in Ba2+.
Observed reduction in excitability is essentially ohmic and is due to three factors: decreased membrane resistance near RMP (1.5- to 5-fold), raised action potential threshold (2.6 ± 0.5 mV), and membrane hyperpolarization (7 ± 2 mV for all ROMK1 cells relative to precisely age-matched controls). The latter two factors are directly related to the change in resistance, identifying a single factor that is necessary for silencing and that can be used to predict neuronal response to any foreign channel.
The observed results confirm that ROMK1 is functional in the infected neurons studied and that changes in resting excitability are due to a Ba2+-blockable K+ conductance. The predictability of these changes, and the normal firing patterns of infected cells in Ba2+ even after weeks of infection, suggest that nonspecific metabolic effects of the channel are minimal in 15.4 mM K+. As few as 150 open ROMK1 channels can result in silencing of 20 pA, while higher numbers (up to 1,000) lead to neurons that cannot fire in the absence of Ba2+.
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CONCLUSION |
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![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
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What is the future of K+ channels as
silencing agents? Our results confirm previous acute experiments
showing hypoexcitability in response to a foreign
K+ channel, but underline the importance of
long-term expression to identify adverse affects that may occur in
transgenic animals or in gene therapy. A transgenic silencing
experiment may be regulated by means of inducible promoters with
adjustable levels of induction, such as the tetO-CMV promoter
(Huang et al. 1999) and ideally would be fully
reversible. However, the time scale of induction and reversal with even
the best genetic systems is days to weeks (Chen et al.
1998
), several times longer than the life span of neurons
expressing even the lowest levels of ROMK1. This channel therefore does
not provide an alternative to all-or-nothing lesions or knockouts, at
least in adult neurons.
A weak inward rectifier was chosen for these experiments partially because of its expected ability to hyperpolarize neurons with outward current, but the demonstrated minor role of membrane potential in silencing suggests that this type of channel is not the best. A strong inward rectifier may provide all of the silencing with less or none of the K+ loss; experiments to test long-term expression of such channels are therefore the next step.
There are also neuronal subtypes and developmental stages that require
high K+ medium in dissociated culture, and
analogously, high levels of electrical input in vivo: cerebellar
granule cells (Galli et al. 1995) and retinal cells
(Araki et al. 1995
) are prime examples. As long as these
depolarizing conditions exist, such cells may be silenced but not
killed by the presence of ROMK1, as are hippocampal neurons in high
K+. The channel may therefore be a useful tool
for studying the dependence of neuronal migration and differentiation
on excitability. Whether cells that require elevated
K+ are able to survive ROMK1 expression in vitro
may be easily tested before carrying out transgenic experiments. The
increasingly understood link between apoptotic pathways and
K+ loss (Padmanabhan et al. 1999
;
Pike et al. 1996
) may also make ROMK1 an excellent model
of neurodegeneration.
Finally, our electrophysiological results identify the factors
important for electrical silencing and suggest alternative methods of
achieving this goal. K+ may be too intimately
connected with the cell cycle to allow its balance to be altered, but
any ion channel that doubles a neuron's input conductance will be a
significant silencing agent, as long as its reversal potential is more
negative than RMP. An example would be Cl
channels, which play important inhibitory roles in vertebrate and
invertebrate nervous systems.
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APPENDIX A: Effects of high K+ culture conditions |
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A variety of new or up-regulated channels and functional changes were identified in hippocampal neurons in response to chronic application of 15.4 mM K+. Apart from EIRK, these effects do not alter our analysis of ROMK1, as they occur to the same extent in control and ROMK1 neurons. Nevertheless, they are important for situations in which these cells may be exposed to depolarizing conditions.
Synaptogenesis, cation channels, and TTX-insensitive Na+ current
A lack of synaptogenesis due to prolonged elevation of
K+ has been seen in neocortical neurons
(Baker et al. 1991). We observe a similar effect here,
where no postsynaptic potentials or currents are resolvable in high
K+ control cells even after 14-21 dic. An
identical picture occurs in the ROMK1 cells, not reversed by
Ba2+ application (Fig.
A1). The mechanisms responsible for
this synaptic silence may be different in the two cases: in the ROMK1
cells, high-input conductance may shunt synaptic input. However,
effects of ROMK1 on synapses are experimentally inaccessible under
these conditions.
|
The neurons also show the development of at least one
Ba2+-blockable nonspecific or cation current, so
that by 13 dic, Ba2+ wash-in leads to membrane
hyperpolarization. The effect of Ba2+ on neurons
in long-term high K+ is complex and follows at
least two separate time courses (see Fig. 5A): a rapid
hyperpolarization, followed by a much slower increase in membrane
resistance at voltages near threshold (more positive than 60 mV). On
Ba2+ washout, the membrane potential recovers
rapidly, but the resistance remains at this higher value throughout the
time courses observed (1-2 min). There is no apparent difference
between ROMK1-infected cells and controls.
Additionally, both the ROMK1 and high K+ control
neurons show a TTX-insensitive Na+ conductance
that becomes apparent with the application of 500 µM
Ba2+ (Fig. 7, B and D);
this has also been noted in brainstem motor neurons raised in elevated
K+ (Eustache and Gueritaud 1995).
The conductance is not altered by application of
Co2+ and is therefore not
Ca2+ dependent. It is slightly but not
significantly larger in high K+ controls (38 ± 6 pA/pF, n = 4, occurrence in 4/5 cells in TTX) than
in ROMK1 cells (25.4 ± 3.7 pA/pF, n = 9;
occurrence in 9/10 cells in TTX; P = 0.2). This
indicates that it is a response to the elevated
K+, not to ROMK1, and may in fact occur to a
slightly lesser degree in the latter. Nevertheless, ROMK1 cells and not
controls are able to fire spikes in the presence of 1 µM TTX, 1 mM
Co2+, and Ba2+ (Fig.
A2).
|
EIRK up-regulation: Less in ROMK1 neurons
The only adaptive change that differed between ROMK1 and control neurons involved the inwardly rectifying K+ conductance referred to in this paper as EIRK. As noted in the discussion of RMP, Ba2+ blockade of ROMK1 restores infected neurons to a state more like that of low K+ controls than high K+ controls. So is it correct to subtract the high K+ control I-V from that of the ROMK1 cells to quantify expression levels at very negative potentials, or is all current in ROMK1 cells due to ROMK1?
ROMK1-expressing neurons will not necessarily up-regulate EIRK as do
controls, because ROMK1 prevents the hyperexcitability caused by high
K+ that presumably potentiates development of
native inward rectifiers (Fig. A3). It
is possible to distinguish the two types of conductance by their
sensitivity to Ba2+ at 144 mV. This will
provide an estimate of the percentage of Ba2+-blockable conductance that is due to EIRKs
in ROMK1 cells.
|
Examination of the currents at 144 mV (Tables 2 and
A1) reveals that the
Ba2+-blockable current in completely silenced
cells is significantly greater than in all ROMK1 cells, but the
residual (nonblockable) current is not. Further, the percentage of
current that is blockable increases with increasing expression of
ROMK1. This suggests that the fractional block of EIRK differs from
that of ROMK1, and that as ROMK1 expression levels increase, the
percentage of current due to EIRKs decreases proportionately. If the
current at
144 mV in a high K+ control is
assumed to be made up of IEIRK and a
nonblockable part I0, then the
fraction blocked is given by
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![]() |
(A1) |
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(A2) |
|
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APPENDIX B: Two-compartment localization of currents |
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Electrophysiological data from all cultured neurons in our
experiments, regardless of age, could be fit to a semi-empirical model
with five experimentally measured parameters that describe the time
dependence of the current in response to a voltage step V0
![]() |
(B1) |
A current of this form is consistent with a rapidly charging cell body
and distal dendrites that charge more slowly (details in Nadeau
and Lester 2000). The dendrites are separated from the soma by
a resistance RC = 175 ± 25 M
for
all ROMK1 cells, and this resistance is assumed to be a property of the
cell's anatomy that does not change on Ba2+
wash-in. The voltage at the soma as a function of time is then given by
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(B2) |
![]() |
(B3) |
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(B4) |
![]() |
(B5) |
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(B6) |
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ACKNOWLEDGMENTS |
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We thank B. Khakh, G. Greif, J. Pine, and C. Lindensmith for useful suggestions and discussions.
This work was supported by Burroughs-Wellcome and by National Institute of Mental Health Grant MH-49176.
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FOOTNOTES |
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Address for reprint requests: H. Nadeau, Caltech Biology 156-29, 1200 E. California Blvd., Pasadena, CA 91125 (E-mail: nadeau{at}cco.caltech.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 27 March 2000; accepted in final form 4 May 2000.
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REFERENCES |
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