1 Department of Physiology and Biophysics, University of Texas Medical Branch, Galveston, Texas 77555; and 2 Department of Biochemistry, University of Toronto, Toronto, Canada M5S 1A8
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ABSTRACT |
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In a previous
report [T. J. Kleine, A. Gladfelter, P. N. Lewis, and S. A. Lewis.
Am. J. Physiol. 268 (Cell Physiol. 37):
C1114-C1125, 1995], we found that the cationic DNA-binding proteins
histones H4, H1, and H5 caused a voltage-dependent increase in the
transepithelial conductance in rabbit urinary bladder epithelium. In
this study, results from lipid bilayer experiments suggest that
histones H5-H1 and H4 form variably sized conductive units. Purified
fragments of histones H4 and H5 were used to determine the role of
histone tertiary structure in inducing conductance. Isolated COOH- and NH2-terminal tails of histone H4,
which are random coils, were inactive, whereas the central -helical
domain induced a conductance increase. Although the activities of the
central fragment and intact histone H4 were in many ways similar, the
dose-response relationships suggest that the isolated central domain
was much less potent than intact histone H4. This suggests than the
NH2- and COOH-terminal tails are
also important for histone H4 activity. For histone H5, the isolated
globular central domain was inactive. Thus the random-coil
NH2- and COOH-terminal tails are
important for H5 activity as well. These results indicate that histone
molecules interact directly with membrane phospholipids to form a
channel and that protein tertiary structure and the degree of positive charge play an important role in this activity.
tight epithelium; mammalian bladder; toxicity; ion permeability
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INTRODUCTION |
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CATIONIC PROTEINS (proteins with a net positive charge) are known to be cytotoxic to eukaryotic cells as well as to possess antimicrobial properties. Examples of such cationic proteins are numerous and include protamine sulfate, histones, major basic protein (MBP), and eosinophil peroxidase (EPO) (6, 7, 13, 19). Although histones are normally contained within the nucleus of the cell, conditions such as cell death and lysis cause the release of histones from the cell. Purified histones have been found to increase cell membrane permeability to small monovalent cations and anions, and it has been proposed that this increase in membrane permeability leads to cell swelling and ultimately cell lysis (7). Thus the release of histones may be pathologically important in conditions of significant cell death, such as that which occurs during the breakdown of sperm. In this regard, Mendizabal and Naftalin (12) demonstrated that human semen was toxic to rat colonic mucosa, resulting in a focal loss of epithelial cells (i.e., a loss of the local barrier function of the colon). In addition, some patients with diabetes mellitus suffer from retrograde ejaculation (a painful disorder), which results in the delivery of semen into the lumen of the bladder. Given the cytotoxic nature of histones on the bladder epithelium, the pain resulting from retrograde ejaculation might be caused by the loss of bladder barrier function, allowing ready access of urine to underlying sensory neurons.
The membrane conductances induced by histone share a number of properties with protamine, MBP (unpublished observations), and EPO (unpublished observations), including voltage dependence and reversal by calcium, suggesting a common mechanism among these proteins. Thus the effects of histone on membrane permeability are of interest not only because of a potential pathological role of histones on disruption of the barrier function of colonic and bladder epithelia but also as a model for the mechanism of action of other cationic proteins.
In a previous report (7), it was demonstrated that histones H1, H4, and H5 increase apical membrane permeability in rabbit urinary bladder epithelium, which ultimately led to cytotoxicity. The permeabilities were characterized in terms of the dose-response relationship, voltage dependence, ion selectivity, and reversibility. However, it was unclear whether histone was forming a channel in the cell membrane or was instead increasing activity of a native membrane channel.
Other questions posed by this earlier study involve defining the
structures of the histone molecule that participate in activity. Histones H4 and H5 are two similar yet structurally distinct cationic DNA-binding proteins (Table 1). Both have two
random-coil tails that flank a central domain. This central domain of
H4 contains an -helical region that spans amino acids 55-67
(3). A second span from 70-90 has been demonstrated to be
-helical when histone H4 is associated with the nucleosome but not
when purified histone H4 polymerizes in solution (8). In contrast, the
central domain of histone H5 is globular (2). Another structural
difference between histones H4 and H5 is that the COOH-terminal tails
of histone H4 can associate into
-sheets to form
high-molecular-weight aggregates (8). Although both histone H5 and the
isolated globular domain have been shown to self-associate under
certain conditions (11), significant aggregation of intact H5 in
solution does not occur.
In this report, the following questions are addressed. 1) Do histones induce the formation of a channel? and 2) What are the active domain(s) of the histone molecule? The data presented in this study suggest that histones interact with phospholipids to induce the formation of a channel. In addition, the central fragment of histone H4 (amino acids 25-67) is important for channel formation, whereas the random-coil tails are important in potentiating the activity of the channel-forming domains. For histone H5, the isolated central globular domain was inactive, demonstrating the importance of the random-coil tails for the conductive activity of histone H5.
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MATERIALS AND METHODS |
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Purification of Histones and Histone Fragments
Purification of histones H4 (mol wt 11,294) and H5 (mol wt 20,900) has been described previously (8, 9). In brief, histones were separated from chicken erythrocytes by gel filtration using a Bio-Gel P-10 column (150 × 2.7 cm) eluted with 0.02 M HCl. Fractions contained purified H4 and an H5-H1 (4:1) mixture. H5 was separated from H1 by ion-exchange chromatography. Histones were recovered from concentrated column fractions by precipitation with acidified acetone (0.1% HCl) and then washed with acetone and dried under a vacuum.Production of histone H4 fragments has been described previously (8). Intact H4 was cleaved at three aspartic acid residues (amino acids 24, 68, and 85) as follows. Briefly, H4 was dissolved in 0.25 M acetic acid and heated for 6 h at 105°C. The mixture was then fractionated on a Sephadex G-50 column (150 × 2.7 cm) and eluted with 0.02 M HCl. Eluted fractions were monitored at 200 nm and were appropriately pooled. The pooled fractions were then dialyzed against absolute ethanol, and 10 volumes of acidified acetone (0.1% HCl) were then added to precipitate the peptides. The peptides were washed with dry acetone and dried under a vacuum. Fragments 1-23 and 69-102 were separated by preparative electrophoresis (20) using a Sephadex G-10 column (50 × 1.1 cm) with 0.05 M acetic acid buffer at 500 V (1.4 mA) for 3.5 h.
Purification of the globular domain of H5 has been previously described (2). First, histone H5 (20 mg/ml) was dissolved in 0.2 M K2SO4 and 50 mM tris(hydroxymethyl)aminomethane · HCl buffer, at pH 8. Next, trypsin was added at an enzyme-to-substrate ratio of 1:1,000 at 20°C for 2 h. The digestion was then quenched using 0.02% 1-chloro-3-tosylamido-7-amino-heptanone, and the globular H5 was isolated using a Sephadex G-50 F column. The sample was then dialyzed against 20 mM HCl and recovered by acetone precipitation. All histones were dissolved in distilled, deionized water to make concentrated stock solutions, which were stored at 0°C.
Bilayer Experiments
All phospholipids were purchased in chloroform from Avanti Polar Lipids (Alabaster, AL). The chloroform was evaporated with nitrogen gas, and the lipids were resuspended in decane. Bilayer formation has been described previously (5). The lipids were painted across a 100-200 µM aperture of a Delrin cup until a bilayer with a capacitance of 200-500 pF was formed. Experiments were performed in symmetric 150 mM KCl in twice-distilled water at room temperature. Histone was added to the solution in the cis-chamber, and the trans-chamber was defined as a ground. The solutions in both chambers were stirred with magnetic stirring bars.Voltage was passed, and the resulting current (I) was measured via Ag-AgCl electrodes connected to a bilayer voltage clamp (5). These were continuously monitored on an oscilloscope as well as passed through a pulse-code modulator (Sony) and recorded on videotape. The data were subsequently digitized and analyzed using pClamp 6.0 (Axon Instruments).
Transepithelial Voltage Clamping Experiments
Tissue preparation. Urinary bladders were excised from 3-kg male New Zealand White rabbits and were washed in NaCl Ringer (see Solutions below). The smooth muscle was dissected away, and the epithelium was mounted on a ring of 2 cm2 exposed area. The ring was transferred to a temperature-controlled, modified Ussing chamber (10) where the serosal side of the epithelium was held against a nylon mesh by a slight excess of solution in the mucosal chamber. Both the mucosal and serosal chambers initially held a bathing solution of NaCl Ringer and were aerated with 95% O2-5% CO2 while integral water jackets maintained the temperature of the bathing solution at 37°C. The mucosal chamber was modified to reduce the volume to 4.5 ml (the serosal chamber volume was 15 ml). This was done to minimize the amount of protein used in the experiments. The serosal chamber was stirred by a magnetic spin bar at the bottom of the chamber while the mucosal chamber was stirred by adding the 95% O2-5% CO2 at the bottom of the chamber and allowing it to bubble upward.
Solutions. NaCl Ringer contains (in mM) 111.2 NaCl, 25 NaHCO3, 10 glucose, 5.8 KCl, 2.0 CaCl2, 1.2 KH2PO4, and 1.2 MgSO4. In KCl Ringer, all Na+ salts were substituted with the appropriate K+ salts. Unless otherwise noted, all experiments with histones and histone fragments were performed using KCl Ringer as the mucosal bathing solution. Histones, histone fragments, and amino acid heteropolymers were suspended in distilled H2O as a stock solution that was added in microliter quantities to the mucosal solution. Poly(Lys-Ala) 1:1 and poly(D-Glu-D-Lys) 6:4 were purchased from Sigma (St. Louis, MO).
Transepithelial Electrophysiological Methods
Electrical measurements. All electrical measurements were made under voltage clamp conditions unless otherwise noted. The transepithelial voltage (Vt) was measured with Ag-AgCl wires placed adjacent to either side of the epithelium (serosal solution ground) while I was passed from Ag-AgCl electrodes placed in the rear of each hemichamber. Both sets of electrodes were connected to an automatic voltage clamp (Warner Instruments). Transepithelial resistance and its inverse, transepithelial conductance (Gt), were calculated using Ohm's law from I required to clamp the epithelium 10 mV from the holding voltage under voltage clamp conditions.
Data acquisition. I and voltage outputs of the voltage clamp were connected to an analog-to-digital converter (Axon Instruments) interfaced with a computer that calculated values for resistance and short-circuit current (Isc). Vt and I were continuously monitored on an oscilloscope. All data were printed out with the time of data acquisition and were additionally stored on hard disk.
Equivalent circuit analysis. The method of Yonath and Civan (21) was used to differentiate between an increase in the conductance of the cell membrane or tight junctions. Gt (µS/cm2) was plotted as a function of Isc (in µA/cm2) when Vt = 0 mV in the presence of added protein. This plot was then fit by the equation
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(1) |
Current-voltage relationship.
The steady-state difference current-voltage
(I-V) relationship of the
protein-induced conductance was calculated using the method of Tzan et
al. (19). This method involves measuring the transepithelial
I-V relationships in both the absence
and the presence of added protein; the difference between these two
relationships is the voltage dependence of
I flowing through the protein-induced conductance. First, the tissue was voltage clamped to
Vt = 0 mV, and
the transepithelial I responses to
computer-generated voltage pulses 30 ms long and of increasing
magnitude and alternating polarity were measured. Next, the
transepithelial potential was voltage clamped to 70 mV, protein
was added to the mucosal solution and equilibrated for 3 min, and then
the transepithelial potential was clamped to 0 mV. The conductance was
allowed to reach a steady state before the
I-V relationship was again measured.
The difference between the I-V
relationships in the presence and absence of added protein was then fit
by the constant-field equation to determine the relative ionic
permeabilities of the protein-induced conductance.
Data analysis and statistics. Curve fitting was done on an IBM-AT using NFIT (Island Products, Galveston, TX). Statistics were calculated using INSTAT (GraphPAD Software, San Diego, CA). Data are shown as means ± SE.
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RESULTS |
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In this section, we first report the effects of purified histones on phospholipid bilayers. Then the fragments of histones H4 and H5 and their ability to induce an increase in Gt in rabbit urinary bladder epithelium are compared. Finally, the synthetic proteins poly(Lys-Ala) and poly(Glu-Lys) are tested to determine the role of negatively-charged amino acids.
Histones Induce a Conductance in Bilayers
In previous reports of the conductive effect of histones on rabbit urinary bladder epithelium, it was unclear whether histones were directly inducing channel formation or whether they were indirectly affecting epithelial conductance, for example, by activating a native membrane channel or a second messenger system which would in turn cause an increase in conductance (7). Therefore, purified histones were tested on phospholipid bilayers.First, a 4:1 mixture of histones H5 and H1 (H5-H1) was tested on
phospholipid bilayers. Histone H5-H1 (185 nM) was added to the solution
bathing the cis-side of phospholipid
bilayers composed of a 5:3:2 ratio of phosphatidylserine (PS),
phosphatidylethanolamine (PE), and phosphatidylcholine (PC) (wt/wt/wt).
Voltage was first held at 0 mV
(trans-side ground) during an
equilibration period of several minutes and then was clamped to either
100 mV or 100 mV. At either voltage, channels of variable sizes
appeared (Fig. 1A).
When the voltage polarity was reversed, channels were still evident.
The all points histogram of the I
tracing demonstrates the variability in the magnitude of the channels
(Fig. 1B). In eight bilayers, the
channel conductances ranged in size from 4 to 20 pS. H5-H1 (185 nM) was
also tested on bilayers composed of only PE. Channel activity was
observed at both +100 mV and
100 mV (data not shown). However,
in contrast to the PS:PE:PC bilayers, only single-channel events with a
low probability of opening (0.036 ± 0.005, n = 4) were observed in PE bilayers.
This suggests that histones have a stronger affinity for
negatively-charged phospholipids, which results in increased channel
activity.
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Histone H4 (89 nM) was also added to bilayers composed of a 5:3:2 ratio
of PS:PE:PC. Channels of variable sizes appeared at both 100 mV and
100 mV (Fig.
2A). In
addition to distinct channel openings and closings, there were sporadic
increases in noise that could have been the result of channels
flickering open and closed. Because of this noisy channel activity, the
peaks in the all points histograms were broad and not well resolved
(Fig. 2B). Therefore,
I amplitudes were determined by
measuring each distinct individual opening. The distinct channels were
variable in size, ranging from 2 to 15 pS (53 distinct openings,
n = 2). In another bilayer, after
histone addition, several large spikes (100-160 pS) appeared
followed by breakdown of the bilayer (not shown). These results suggest
that histone H4, like histone H5-H1, is capable of forming channels of
variable sizes in phospholipid bilayers.
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Activity of the Histone Fragments
The activity of whole histones has previously been described in rabbit urinary bladder epithelium (7). In this report, the effects of the histone fragments are tested to identify the active domains. The activity of the fragments is characterized in terms of time course, dose response, voltage sensitivity, ion selectivity, and site of action. Differences between the activity of the histone fragments and the intact histones are also described. Unless otherwise noted, the mucosal solution was a KCl Ringer in all histone and histone fragment experiments.The effect of histone fragments on
Gt of rabbit urinary
bladder epithelium.
A previous report suggested that histones induce a voltage-sensitive
conductance in the apical membrane of rabbit urinary bladder (7). In
urinary bladder epithelium, the apical membrane has a high resistance
to ion flux, whereas the basolateral membrane is quite permeable to
K+ and
Cl. Therefore, the voltage
across the apical membrane can be controlled by the transepithelial
potential. The apical membrane voltage (Va) is
calculated as the difference between
Vt and the
basolateral membrane voltage
(Vb), which is
relatively constant at
55 mV [as previously determined
using microelectrodes (10)]. For example, when
Vt is clamped at
70 mV (serosa ground),
Vb is
55
mV, and therefore
Va is +15 mV
relative to the mucosal solution. If
Vt is clamped to
0 mV, the apical membrane cell potential is
55 mV. Under normal
conditions, Gt of
the rabbit urinary bladder epithelium displayed little or no voltage
sensitivity. However, when either histone H1, H4, or H5 was added to
the mucosal solution, there was a rapid, voltage-sensitive increase in
Gt when
Vt was clamped
from
70 mV to 0 mV (7). All three of the histones induced an
increase in Gt
only when the voltage gradient across the apical membrane was cell
interior negative.
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(2) |
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Dose-response relationship. The dose-response relationship also indicated that the histone fragment H4-(25-67) was much less potent than intact H4. The magnitude of the induced conductance was determined as a function of concentration for both the fragments and intact histone H4 (Fig. 4). The H4 fragments (1-23), (69-102), and (86-102) did not alter Gt at any of the tested concentrations. The globular domain of H5 also did not elicit a response for concentrations ranging from 102 to 3,700 nM (3 tissues, data not shown). In contrast, the magnitude of the conductance increased as a function of protein concentration for both H4-(25-67) and intact histone H4. The shape of the dose-response relationship for the H4-(25-67) fragment is sigmoidal. Such a sigmoidal relationship possibly suggests that several molecules of the fragment combine to induce a conductance. This is in contrast to the dose response of intact histone H4, which is hyperbolic.
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Voltage sensitivity. Both the magnitude and the rate constant of the H4-(25-67)-induced conductance increased as a function of the applied voltage. In addition, the H4 fragment was more sensitive to the applied voltage than intact histone H4. After protein addition and equilibration, the tissue was clamped to various Vt and the magnitude and rate constant were measured at each voltage. As shown in Fig. 5, A and B, both the magnitude and the rate constant were exponential functions of the transepithelial (and apical) membrane voltage. Values were normalized to the conductance change or rate constant measured at 0 mV to correct for tissue variability.
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(3) |
Site of action.
Previously, it has been shown that intact histone H4 predominantly
affects the apical, rather than junctional or basolateral, membrane
permeability. To determine if the central helical domain has the same
site of action, the method of Yonath and Civan (21) was used (see
MATERIALS AND METHODS). While
Vt was clamped to 0 mV, the changes in conductance
(Gt) and
Isc were
monitored. Changes in
Gt can occur at
three sites: the apical membrane, basolateral membrane, or tight
junctions. If the protein-induced conductance increases as a linear
function of Isc
(see Eq. 1), this suggests that only
the cellular conductance (apical and/or basolateral membrane
conductance) was increased by histones. The best fit values to
Eq. 1 for the fragment are
Ec = 60 ± 3 mV and
Gj= 28 ± 6 µS/cm2
(n = 6). As shown in Fig.
6, the relationship between the change in
Gt and the change
in Isc was linear
for the
-helical fragment, with a slope similar to that of intact
histone H4. This suggests that the fragment acts on the apical membrane
and that removal of the tails does not alter the specificity of the
site of action.
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Ion selectivity.
The H4-(25-67)-induced conductance was nonselective for
K+ and
Cl. The ion selectivity was
determined from the steady-state difference I-V relationship (see
MATERIALS AND METHODS). An example
of this relationship, which was linear, is shown in Fig.
7. The data were fit by the constant-field
equation to determine the K+ and
Cl
permeabilities. For 426 nM H4-(25-67), the best fit value for the
K+ permeability was 3.6 ± 1.2 × 10
8 cm/s
(n = 4), and the ratio of
Cl
to
K+ permeability
(PCl/PK)
was 0.8 ± 0.2 (n = 4), indicating
that the fragment-induced conductance was nonselective for these two ions. This is in agreement with the
I-V relationship for intact histone H4
and suggests that the ion specificity of the induced conductance is
conferred by the central helical domain.
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Reversibility of the induced conductance.
Previously, it has been demonstrated that the intact histone H4-induced
conductance can be reversed be changing
Vt from 0 mV back
to 70 mV (7). To determine if the loss of the
NH2- and COOH-terminal tails
affected voltage-dependent reversal,
Vt was clamped
back to
70 mV after the fragment-induced conductance reached a
plateau at 0 mV. The time course of the conductance reversal is shown
in Fig. 8. Note that the conductance
increased first before decreasing. This response was observed in all of the voltage reversal experiments (n = 20). The average increase was 31 ± 3 µS/cm2. In contrast, only 44%
of the intact histone H4 voltage reversals were reported to show this
response (7). This initial jump was hypothesized to result from the
intact histone H4 partitioning through the cell membrane into the
cytoplasm, where it would induce a conductance when the voltage
gradient across the apical membrane was cell interior positive. This
was further supported by the serosal addition of histone resulting in
an increase in apical membrane conductance, with the opposite voltage
polarity as mucosal histone, suggesting that histone entered the cell
through the basolateral membrane rather than by crossing the tight
junctions and entering into the mucosal solution (7). Because the
initial jump was more frequent with the H4-(25-67) than with
intact histone H4, this suggests that the fragment may partition
through the membrane at
Vt = 0 mV more
easily than intact histone H4. After the initial jump, when clamping
from 0 mV to
70 mV, the reversal of the H4-(25-67)-induced
conductance followed the form of a double exponential, consistent with
intact histone H4.
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Role of Acidic Amino Acids
As the fragment experiments demonstrate, the tertiary structure of histones is important for their conductive activity. It has been demonstrated that positively charged amino acids located within the protein molecule are important as well (18). However, the role of negatively charged amino acids within the protein molecule has not been characterized. To determine the role of acidic amino acids within the protein, two synthetic molecules were tested: poly(Lys-Ala) 1:1 and poly(D-Glu-D-Lys) 5.7:4.3. Both of these proteins had a molecular weight that ranged from 20,000 to 50,000; the average molecular weight was 41,600 for poly(Lys-Ala) and 23,000 for poly(Glu-Lys), as determined by viscosity measurements (Sigma). These proteins are similar in that approximately one-half of each protein is composed of cationic amino acids. However, they differ in their net charge density. The charge density is defined as the percentage of the total protein molecule that is composed of similarly charged amino acids (18). The net charge density is the difference between the positive and negative charge densities of the protein. Poly(Lys-Ala) had a net charge density of 50% positive charge, whereas poly(Glu-Lys) had a net 14% negative charge density.The effect of poly(Lys-Ala) and poly(Glu-Lys) on
Gt.
Each synthetic protein was tested on the urinary bladder epithelium in
the same manner as the histone fragments; protein was added to the
mucosal solution while
Vt was clamped at
70 mV and equilibrated for 3 min. Then
Vt was clamped to
0 mV. Typical time courses for each protein are shown in Fig.
9. Note that the concentration of
poly(Glu-Lys) was 24-fold higher than poly(Lys-Ala).
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(4) |
The dose-response relationships. Poly(Lys-Ala) was much more potent than poly(Glu-Lys), as indicated by the dose-response relationship. The relationship between the magnitude of the conductance change and the concentration of added protein was determined in the same manner as described for the histone fragment (see above). For poly(Lys-Ala), the time courses were fit by Eq. 4, and Gp1 and Gp2 were added to determine the total conductance change. As shown in Fig. 10, the magnitude of the conductance induced by either protein increased as a function of protein concentration, but poly(Lys-Ala) was much more potent than poly(Glu-Lys). There are several possibilities to explain the reduced activity by poly(Glu-Lys). One possibility is that the net negative charge repels poly(Glu-Lys) from an anionic binding site for cationic proteins. Another explanation is that, by interacting electrostatically, the acidic residue glutamic acid neutralizes the basic amino acid lysine, which could result in loss of the voltage sensitivity of the peptide. An alternative explanation is that the poly(Glu-Lys) molecules electrostatically interact to form aggregates that are inactive.
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DISCUSSION |
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In this paper, the components of both the cellular membrane and the protein that are important for conductive activity were examined. First, the identity of the membrane-binding site and the ability of histone to induce ion channel formation were explored using phospholipid bilayers. The results suggest that histone is capable of inducing channel formation and that this activity is enhanced by the presence of anionic phospholipids in the membrane. The importance of histone tertiary structure and the effect of negatively charged amino acids within the protein molecule were explored in an effort to elucidate the structural features of the protein that contribute to activity.
Histone Channels and Membrane-Binding Sites
The results of the phospholipid bilayer experiments are important from two perspectives: first, because channel activity occurred in the presence of histone, this indicates that histone is capable of forming channels; second, the channel activity was increased in anionic phospholipid-containing bilayers compared with bilayers composed only of neutral phospholipids. This suggests that anionic membrane-binding sites, although not required for histone activity, enhance channel formation. This is perhaps a result of the electrostatic interaction between histones and the anionic phospholipid, resulting in an increased amount of histone binding or greater stability of the histone-phospholipid interaction. Histones have been reported to preferentially bind to anionic phospholipids (16). Purified histones were demonstrated to bind to the anionic phospholipids cardiolipin and PS with high avidity but not to the zwitterionic phospholipid PC.Histone Fragments
The 25-67 fragment of histone H4 was capable of increasing Gt in a manner similar to intact histone H4. Both the fragment and H4 conductances displayed comparable time courses, sites of action, voltage dependence, and ion selectivity. Because of the limited quantity of fragment available, the long-term toxicity was not determined. Therefore, it is yet to be determined if the fragment is able to induce the same degree of toxicity as intact histone H4.The conductive activity of the H4-(25-67) fragment suggests that
this fragment is important for channel formation and may contain the
channel-forming domain. Both the structure of this fragment and of
intact histone H4 has been described in detail (3, 8). Comparison of
the structures may help explain the similarities and differences in the
activities of these two proteins. With the use of fragments of histone
H4, it has been determined that the stretch of amino acids from 50 to
67 is critically involved in the formation of both the -helix and
multimeric aggregates. The
-helix is composed of two sections,
residues 55-67 and 70-90. The helical wheel diagram of the
helical portions of histone H4 suggests that the stretch of amino acids
involved in helix formation and aggregation (amino acids 50-67) is
somewhat amphipathic in character (Fig.
11). The hydrophilic amino acids are
located on one side of the helix, opposite to the majority of the
hydrophobic amino acids, making one side of the helix much more polar
than the rest of the helix. This suggests that histone H4 [and
the H4-(25-67) fragment] may belong to a family of
amphipathic
-helical proteins that increase membrane permeability
(4, 14, 15, 22). These proteins are believed to aggregate into
barrel-like structures, with their outward-facing hydrophobic sides
interacting with the phospholipid bilayer, while the hydrophilic faces
line the channel. Further investigation is necessary to determine if histone H4 behaves similarly.
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When the H4-(25-67) fragment aggregates in solution, the portion of the H4-(25-67) fragment that is not helical (residues 25-54) is incorporated into the aggregates rather than being free in solution. In contrast, histone H4 has an additional stretch of NH2-terminal tail (residues 1-24) that is highly cationic and is a random coil in solution. The carboxyl tail of histone H4 is also a random coil but is not quite as cationic as the amino tail; the COOH-terminal tail is important for the formation of high-molecular weight aggregates by histone H4 (23).
Both histone H4 and the 25-67 fragment are each long enough to
span the cell membrane (>20 amino acids); therefore, individual protein molecules may form the conductive unit. In addition, both of
these proteins aggregate, and therefore channels might also be formed
by the polymerized proteins. Previous reports by Lewis et al. (8)
indicate that these two proteins aggregate differently. Both intact
histone H4 and H4-(25-67) rapidly aggregate along the -helical
residues in a parallel fashion. Intact histone H4 then forms
high-molecular weight aggregates by slowly forming
-sheets along the
COOH-terminal tails. Fragment H4-(25-67), in contrast, quickly
forms
-sheets at the
NH2-terminal of the fragment (amino acids 25-34). The fragment, however, is not able to form high-molecular weight aggregates, because the COOH-terminal is required
for this process (23).
There are a number of possible explanations for the lower potency of
the H4-(25-67) fragment. 1) The
-helical region from amino acids 70-90, which is found in
intact histone H4 but not in the fragment, may contribute to the
formation of a more stable membrane channel. Studies suggest that
increasing the length of an amphipathic
-helix increases both
channel formation in bilayers and toxicity in bacteria (1).
2) The
NH2- and/or the
COOH-terminal tails of histone H4 are important for membrane binding or
for stabilizing the conductance in the membrane. Loss of the tails would then either make it more difficult for the fragment to associate with the membrane or result in the fragment dissociating from the
membrane more easily. 3) The
-structure of the fragment aggregates may impede their ability to
form a conductance. 4) The
aggregation of the COOH-terminal tails of intact histone H4 into
high-molecular-weight aggregates could lead to the formation of larger
channels than H4-(25-67), which does not form high-molecular
weight aggregates. 5) The
fragment has a weaker voltage sensitivity than intact H4. Preliminary
analysis of washout experiments suggests that the rate constants at
which both histone H4 and the fragment become nonconductive at 0 mV are
similar, whereas the rate constant of formation of the conductance is
much slower for the fragment compared with intact H4. This suggests
that the difference in the dose-response relationships (i.e., the
maximum conductance and binding affinity) is because of the fragment's
slower rate constant for the formation of a conductance than H4.
Interestingly, the central domain of histone H5 did not induce a
conductance. One possible explanation is that the positively charged
amino acids within this central domain are not accessible to the
membrane surface and thus cannot associate with the cell membrane
binding site. The three-dimensional structures of both histone H5 and
the isolated globular domain have been described previously (2, 17).
The central domain is compact and is composed of three -helical
regions and an anti-parallel
-sheet. This complex structure may not
be able to interact with the cell membrane because of steric hindrance.
The COOH-terminal of histone H5 is a long and highly charged
random-coil tail. This tail is composed of amino acids 103-190 and
is 50% cationic. In contrast, the
NH2-tail is small; it spans
residues 1-21 and is 29% cationic. These random-coil tails, which
have been removed from the purified globular domain, may be necessary
for membrane binding of histone H5. Both histone H5 and its globular
domain have been reported to form high-molecular weight aggregates
(11). For globular H5, multimers ranged from 2 to 14 monomers. It is as
yet unclear whether individual protein molecules or aggregates are
responsible for the conductive activity of histone H5.
It has been reported that histone H1 also increases membrane permeability in rabbit urinary bladder epithelium (7). Histone H1 is structurally very similar to histone H5, and histone H5 is regarded as an "extreme variant" of histone H1 (17). They have similar molecular weights (21,000 for H5, 24,000 for H1) and similar structures. Both have a central globular domain flanked by random-coil tails (2). The major structural differences between these two proteins are that histone H5 contains more arginine and serine than histone H1 and that histone H5 has a shorter NH2-terminal tail (21 amino acids compared with 35 amino acids for H1). Histones H1 and H5 display another interesting difference; the globular domain of histone H1 does not form aggregates as readily as the globular domain of histone H5 (11). One might predict that, because of the structural similarities between H1 and H5, the isolated central domain of histone H1 would be inactive.
Predicted Activity of the Histone Fragments Based on Positive Charge
The model developed by Tzan et al. (18) is useful in predicting the relationship between cationic charge and induced conductance. The conductance induced by cationic proteins increased as the square of both the total number of cationic residues and the density of the cationic charge within the protein molecule. This model was developed using synthetic proteins that are composed of cationic and neutral amino acids and are random coils in solution. With the use of this model and histone H1 as a reference molecule, the magnitude of the conductance change that would be induced solely on the basis of positive charge can be predicted for the histone fragments. The calculated values for the fragments indicate that they would be predicted to induce an appreciable change in conductance at the concentrations used in the experiments. There are a number of possible explanations for the lack of activity that was demonstrated by these molecules. Some of the fragments may be too small to be active. For example, the smallest fragment was 86-102, which is only 16 amino acids long and therefore is not long enough to span the cell membrane. The fragment 1-23 is predicted to be the most active of the fragments based on charge but is barely long enough to span the membrane and may not be able to form a stable conductance.Acidic Amino Acids
Addition of negatively charged molecules such as DNA or pentosan polysulfate has been demonstrated to decrease the conductive effect of histone (7), most likely by an electrostatic interaction that neutralizes the cationic charge of the histone. These data suggest that negative charges within the protein molecule also are inhibitory, although the mechanism of inhibition is unclear.The differences in the magnitude of the conductance changes induced by poly(Lys-Ala) and poly(Glu-Lys) are not a result of the difference in positive charge between the two molecules. The model developed by Tzan et al. (18) was also used to predict the magnitude of the conductance change that would be induced by both poly(Lys-Ala) and poly(Glu-Lys). Poly(Lys-Ala) is about as active as predicted, whereas poly(Glu-Lys) is much less active than predicted on the basis of its positive charge. This deviation by poly(Glu-Lys) suggests that the negative charge is reducing the ability to induce a conductance. However, the ability to induce a conductance was not entirely abolished by the net negative charge of the molecule. One possible explanation is that because the synthetic proteins are made by a random distribution of amino acids, they are therefore a heterogeneous mixture of proteins with an average ratio of 6:4 glutamic acid to lysine. A certain percentage of the poly(Glu-Lys) will have a higher concentration of positive charge (and an equivalent amount will have a lower proportion of positive charge). The portion with the higher degree of cationic charge might have a sufficient amount of positive charge so that the net charge on the protein molecule is positive. A net positive charge might then result in the protein being able to induce a conductance.
Summary
These results indicate that histone is capable of forming a conductive unit and identify the central portion of histone H4 as the domain that is responsible for many of the conductive properties of this molecule. The fragment of histone H4 that spans amino acids 25-67 forms a voltage-dependent, non-ion-selective conductance in the apical membrane of rabbit urinary bladder epithelium. This region is predominantly ![]() |
ACKNOWLEDGEMENTS |
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This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant DK-51382 to S. A. Lewis, Medical Research Council of Canada Grant MT-5453 to P. N. Lewis, and James W. McLaughlin Fellowship Fund to T. J. Kleine.
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FOOTNOTES |
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Address for reprint requests: S. A. Lewis, Dept. of Physiology and Biophysics, Univ. of Texas Medical Branch, 301 Univ. Blvd., Galveston, TX, 77555-0641.
Received 20 June 1997; accepted in final form 19 August 1997.
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