From the Program in Molecular Pharmacology and
Therapeutics, Memorial Sloan-Kettering Cancer Center, New York
10021 and the § Department of Chemistry and Lombardi Cancer
Center Program in Tumor Biology Georgetown University,
Washington, D. C. 20057
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ABSTRACT |
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In previous work (Weisburg, J. H., Curcio,
M., Caron, P. C., Raghi, G., Mechetner, E. B., Roepe, P. D., and Scheinberg, D. A. (1996) J. Exp. Med. 183, 2699-2704), we showed that multidrug resistance (MDR) cells created by
continuous selection with the vinca alkaloid vincristine (HL60
RV+) or by retroviral infection (K562/human MDR 1 cells)
exhibited significant resistance to complement-mediated cytotoxicity
(CMC). This resistance was due to the presence of overexpressed
P-glycoprotein (P-GP). In this paper, we probe the molecular mechanism
of this phenomenon. We test whether the significant elevated
intracellular pH (pHi) that accompanies P-GP overexpression is
sufficient to confer resistance to CMC and whether this resistance is
related to effects on complement function in the cell membrane. Control HL60 cells not expressing P-GP, but comparably elevated in cytosolic pHi by two independent methods (CO2
"conditioning" or isotonic Cl A tumor multidrug resistance
(MDR)1 phenotype is
frequently (but not always) due to the overexpression of the
mdr1 gene product, a 170-180-kDa glycoprotein known as
P-glycoprotein (P-GP) or human MDR 1 protein (1). A careful distinction
should be made between the MDR phenotype(s) observed in cells exposed
to or selected with various chemotherapeutic drugs versus
the phenotype mediated solely by P-GP overexpression (2). One
hypothesis proposed to explain the contribution of P-GP to MDR is the
"drug pump" model, in which P-GP hydrolyzes ATP to actively efflux
a variety of chemotherapeutic drugs such as anthracyclines, vinca
alkaloids, and epipodophyllotoxins out of the cell against a
concentration gradient (1, 3). Although this model accounts for some of the properties of MDR cells, it fails to explain resistance to complement-mediated cytotoxicity (CMC) observed in some MDR cells (4)
and other phenomena (reviewed in Ref. 5).
Many investigators working to improve cancer therapy have proposed that
immunologic approaches to killing cancer cells (such as CMC) might
circumvent chemotherapeutic drug resistance. One of the assumptions of
an immunological approach to the treatment of cancer is that cells that
are resistant to cytotoxic drugs (e.g. MDR tumor cells)
should not be cross-resistant to immunotherapy, since the mechanisms of
cytotoxicity are so widely different. Because complement mediates
cytotoxicity via mechanisms directed to the outside surface of the
cell, the protective functions of P-GP overexpression in MDR tumor
cells should not be expected to be relevant to conferring resistance to
CMC; i.e., antibody molecule juxtapositioning following
binding to specific antigen initiates the complement cascade,
culminating in the formation of the membrane attack complex (MAC). When
MACs are inserted into the plasma membrane of the cell, nonspecific
pores as large as 100 Å in diameter are formed, causing cell lysis
(6). We showed previously, however, that a MDR HL60 variant (HL60
RV+), selected on vincristine that also overexpresses P-GP
is resistant to CMC relative to the parental HL60 myeloid leukemia
cells. Differences in killing were independent of antibody isotype
(either IgG and IgM) and target antigen. Resistance to CMC was also
observed in K562 cells overexpressing P-GP via retroviral infection and
not preselected with chemotherapeutic drugs. The immunological
resistance in both MDR cell lines was reversed by the calcium channel
blocker verapamil and the F(ab')2 fragments of the
P-GP-inactivating monoclonal antibody UIC2. These data are consistent
with P-GP overexpression mediating resistance to CMC. Elucidating the
molecular mechanism of this resistance phenomenon is important, because
frequently invoked models for how P-GP contributes to MDR
(e.g. the "drug pump" models) do not provide a framework
for explaining P-GP-mediated resistance to CMC.
MDR cells typically have elevated intracellular pH (pHi) and
decreased plasma membrane potential (Vm) (7-12), and this is apparently due to P-GP overexpression and not exposure to
chemotherapeutic drugs (2). Hence, another model for P-GP's action
(frequently referred to as the altered partitioning model) suggests
that alterations in pHi and/or Vm that accompany the overexpression of P-GP indirectly affect partitioning of
chemotherapeutic drugs, but P-GP does not directly pump them (5, 9, 11,
12). This model may offer a more reasonable framework for addressing
resistance to CMC in MDR cells.
Higgins, Sepulveda, and colleagues (13-15) have suggested that P-GP
may translocate Cl Thus, in this study we examined complement resistance in the
HL60/RV+ system and its relationship to pHi
perturbations. Two independent approaches (Cl The final step of the complement cascade is formation of MACs, which
are large pores in the plasma membrane of the cell that destroy osmotic
balance and lead to cell lysis. MACs consist of five interacting
proteins (C5b, C6, C7, C8, and C9) that, when sequentially activated
upon antibody binding to the target cell, expose hydrophobic residues
and insert into the membrane. Association of these five proteins within
the membrane leads to formation of nonspecific trans-membraneous
channels (19, 20). "Sublytic" dilute concentrations of complement
proteins may form precursors of the full MAC (e.g.
"partial MACs") that are more specific with regard to transport
properties and perhaps even voltage-regulated (21). The pore diameter
of the completely assembled MAC is variable and is essentially
determined by the number of C9 molecules that contribute to pore
formation. MAC stoichiometry is described as (C5b,C6,C7,C8)1C9n, where n = 9-18.
We measured the formation of functional pores with different
characteristics by continuous monitoring of fluorescence methods (9)
and determined the molecular stoichiometry of MAC structure by
purification of MAC complexes harboring radiolabeled subunits. The data
suggest that increased pHi reduces the rate of formation of
functional "full MACs" (i.e.
(C5b-C8)1(C9)n > 9 complexes) in the
cell membrane of target cells.
The results have important consequences for better understanding the
role of P-GP overexpression and chemotherapeutic drug exposure in tumor
resistance phenomena. They also define an important biochemical
parameter for efficient manipulation of immune mediated killing and the
study of the terminal complement cascade.
Materials--
2',7'-Bis(carboxyethyl)-5,6-carboxyfluorescein
acetoxymethyl ester (BCECF-AM) and nigericin were purchased from
Molecular Probes, Inc. (Eugene, OR); H2DIDS was purchased
from Sigma; and baby rabbit complement was purchased from Pel Freez
(Brown Deer, WI). All were used without additional purification.
Cell-Tak was from Becton Dickenson and was used as described by the
manufacturer. Human complement (plasma) was obtained from normal donors.
Cells and Antibodies--
HL60 cells (acute myeloid leukemia,
CD15+, and CD33+) were maintained at Memorial Sloan-Kettering Cancer
Center. HL60/RV+ (a P-GP-overexpressing MDR HL60 variant
selected by continuous exposure to vincristine) was the generous gift
of Dr. Melvin Center (Kansas State University, Manhattan, KS) and was
provided by Dr. Ellin Berman (Memorial Hospital). RV+ cells
were grown in RPMI plus 10% fetal calf serum at 37 °C in an
atmosphere of 5% CO2 and in the presence of 120 mM (0.1 mg/ml) vincristine (Lilly) to maintain the MDR
phenotype. These cells were cultured without vincristine for one
passage prior to experiments. This practice did not reduce MDR or the
level of P-GP expression.
The humanized monoclonal antibody M195 was prepared as described (22,
23), and M31, a murine anti-CD15 antibody, was established at Memorial
Sloan-Kettering Cancer
Center.2 Rabbit anti-human
anti-C5b-9 antibody was purchased form Calbiochem (La Jolla, CA).
Single Cell Photometry and Manipulation of pHi--
As
described previously (10, 16, 24), we have constructed a single cell
photometry apparatus using a Nikon diaphot epifluorescence microscope
and a Photon Technology alpha-scan fluorometer. For BCECF experiments,
a 510-nm dichroic and 530-nm bandpass filter was positioned beneath the
stage, and the excitation monochromator was flipped between 439 and 490 nm by the computer (see Ref. 24 and references therein for additional detail).
Cells (1.0 × 106) were washed and resuspended in 200 µl of a solution consisting of 50 µl RPMI 1640 and 150 µl sterile
filtered HBSS (118 mM NaCl, 5 mM KCl, 24.2 mM NaHCO3, 1.3 mM
CaCl2, 0.5 mM MgCl2, 0.6 mM Na2HPO4, 0.5 mM
KH2PO4, 10 mM glucose). The cells were incubated with 5 µM BCECF-AM for 20 min at 37 °C
and 5% CO2. Next, the cells were adhered to glass
coverslips (Corning Glassworks, 18 mm2/0.11-mm thick) using
Cell Tak (Becton Dickenson) and incubated for an additional 20 min at
37 °C and 5% CO2 and in the continued presence of
BCECF-AM. Coverslips were then mounted in a home-built perfusion
chamber (10) and continuously perfused at a constant rate
(approximately 5 ml/min) with HBSS buffer equilibrated with 5%
CO2 and to 37 °C. Uniform BCECF staining was verified
visually and by monitoring the intensity of 490 nm excitation (535 emission) and was found to be similar between the cell lines.
Excitation was limited to the time of data collection to limit
photobleaching. HBSS was continuously purged with 5% CO2,
and a fine jet of 5% CO2 was directed over the mounted
coverslip to maintain buffer pH, which was monitored with a
microelectrode. Calibration of intracellular BCECF was by the
K+/nigericin titration approach (25), but in single cell
mode (10, 16, 24), wherein HBSS harboring nigericin was continuously flowed over the cells. Steady state pHi was calculated using
the best fit to the calibration data (described in detail in Ref.
2).
To elevate pHi of the control HL60 cells, which do not express
measurable P-GP (8), two approaches were used. First, HL60 cells
underwent elevation of pHi by manipulation of external
CO2 as described (2). In brief, cells were grown at 5%
CO2 in normal RPMI 1640; shifted to a 10% CO2
atmosphere, which causes rapid acidification of the medium; and then
grown in this atmosphere for 2-7 days. The "acid-conditioned"
cells were then placed back in a 5% CO2 atmosphere
in the presence of fresh pre-equilibrated (to 5%
CO2) medium for 1 or 2 h. As described (Ref. 2;
see "Results") this results in an "overshoot" of pHi, since alkalinizing mechanisms induced by growth at 10% CO2
are still functioning (at least for several hours) in the reduced CO2 environment. Detailed analysis of the conditioning
method is presented in Ref. 2.
Alternatively, pHi was elevated by Cl
To examine whether anion exchange was responsible for the alkalization
of the cells under these Cl Complement-mediated Cytotoxicity in HBSS or Cl
To examine if Cl Radiolabeling of HuM195--
Proteins were iodinated using the
Cloramine T method and purified by size exclusion chromatography
(27).
Modulation and Internalization of
125I-HuM195--
HL60 and RV+ cells were
washed and resuspended to a concentration of 1.0 × 106 cells/ml. 125I-HuM195 was added to the
cells to a final concentration of 1 µg/ml. At 0, 0.5, 1.0, 2.0, and
4.0 h, a 200-µl aliquot of cells was removed and washed three
times with ice-cold PBS. Cells were resuspended in 1 ml of stripping
buffer (50 mM glycine, 150 mM NaCl, pH 2.8),
allowed to incubated at room temperature for 10 min, and then
centrifuged at 1500 rpm in a Sorvall GLC 2B centrifuge. Counts/min in
the supernatant fluid (containing cell surface Ig eluted by acid wash)
and the cell pellet (containing internalized Ig) were then quantified
by liquid scintillation spectrophotometry (Packard).
Flow Cytometric Analysis of MAC Formation Kinetics--
1.0 × 107 HL60 or RV+ cells were washed with RPMI
medium. The cell pellet was then resuspended in 800 µl of a 1:10
dilution of M31 antibody and 800 µl of a 1:3 dilution of human serum.
In control tubes, medium was added instead of the human serum. The cells plus antibody and serum were incubated at 37 °C. At 5, 10, 15, 20, and 30 min of incubation, a 160-µl aliquot was removed, and 2 ml
of stripping buffer (see above) was added. This was done to prevent the
secondary antibody (goat anti-mouse fluoroscein isothiocyanate-labeled
antibody) from reacting nonspecifically with the IgM (murine M31
antibody) bound to the cell surface. The cells were incubated with
stripping buffer for 10 min at room temperature and then centrifuged at
750 × g and 4 °C. The cell pellet was resuspended
in 500 µl of ice-cold 2% paraformaldehyde and allowed to incubate on
ice for 15 min in order to fix the MACs. The paraformaldehyde was
removed by washing three times with PBS, and an anti-MAC (C5b-9)
antibody was then added to the cells. This antibody binds to the
neoepitope of C9 that forms during polymerization, but it does not
react with free C9. Anti-C5b-9 antibody was incubated with the cells
for 30 min on ice, unbound antibody was then removed by washing the
cells with PBS, and goat anti-mouse-IgG-fluoroscein isothiocyanate was
added and allowed to bind to primary antibody for 45 min on ice and in
the dark. The cells were then washed with PBS and resuspended in 500 µl of 2% paraformaldehyde. Samples were analyzed on an EPICS-Profile II flow cytometer (Coulter, Hialeah, FL). Cell viability of the samples
collected at 30 min was determined by trypan blue exclusion.
In some experiments, pHi was first altered by CO2
pulse. 1.0 × 107 HL60 cells were
CO2-pulsed for 2 h before examining the kinetics of
MAC formation and were washed in pre-equilibrated RPMI medium at 5%
CO2. The cells were diluted in M31 antibody and human
complement and allowed to incubate at 37 °C as above. As a control,
cells were diluted in antibody and medium without complement but
otherwise subjected to similar treatment. At 5, 10, 15, 20, and 30 min
after the addition of antibody and complement, a 160-µl aliquot was removed from the reaction tube and evaluated as described above.
Radiolabeling Complement Components C7 and C9--
Proteins were
iodinated using the chloramine-T method and purified by size exclusion
chromatography (27). C9 was labeled with 125I; C7 was
labeled with 131I.
Determination of Pore Size and Number of MACs on HL60 Cells under
Cl
The retention time of free radiolabeled C7 and C9 (uncomplexed with
other MAC components) was determined by chromatography on the same
column as above after mixing with carrier human serum. 200 µl of
131I-C7 and 125-I-C9 were added to 600 µl of
undiluted human washed and resuspended in 0.5 ml of RPMI medium.
Photometric Analysis of MAC-mediated Transport--
1.0 × 106 HL60 or RV+ cells were washed and
resuspended in 0.5 ml of RPMI medium. The cells were then transferred
into an Eppendorf tube, and 1 µl of 2.5 mM BCECF-AM in
dry Me2SO was added. The cells were allowed to incubate
with BCECF-AM for 30 min at 37 °C and then washed with HBSS or
Cl
In analyzing these data, we also found that the change in quantum
efficiency of 490-nm excitation is greater than the change at 439 nm
upon movement of the probe from the cytosol to the extracellular space;
thus, exit of the probe from cells in a continuously mixed suspension
also results in a measurable change in the 439/490-nm excitation ratio
(see "Results"). This is probably due to a much greater affect of
solvent polarity on the transition revealed by the 490-nm peak,
relative to the transition revealed at 439 nm. Such an effect
complicates analysis of pHi changes that might occur during an
experiment wherein total BCECF response in a continuously mixed
suspension of cells was being monitored. Analysis of these data to
separate BCECF leak from cytosolic pH changes will be presented
elsewhere.3
BCECF fluorescence data were collected as PTI data files on a PTI
alpha-scan fluorometer, converted to ASCII format, and imported into
QuattroPro 7 software (Corel). To obtain quantitative parameters for
the 439-nm BCECF, data were fit to a function of the form,
CO2 Conditioning Causes Alkalinization of
pHi in HL60 Cells--
One difficulty in accurately
measuring the effects of altered pHi on cellular biochemistry
is the general inability to stably modulate pHi via
nondeleterious means for sufficient periods of time. For example,
chemotherapeutic drug resistance assays require hours to days of
incubation with cytotoxic drugs, but precise manipulation of
pHi via the use of ion transport inhibitors or ionophores is
either toxic, transient, or both; hence, investigating the role of
altered pHi in drug resistance phenomena is difficult and
requires special methods (see Ref. 2). In the present study, we wished
to assess the importance of pHi in CMC, which requires at least
30 min to measure appropriately; thus, we required methods to stably and safely alter pHi within this period of time. As reported previously (2), some cell populations can be manipulated to a higher
average pHi by CO2 conditioning procedures (which should be distinguished from CO2 "pulse" methods that
induce pHi changes on a much shorter time scale; see Ref. 2).
Thus, upon returning HL60 cells to a 5% CO2 atmosphere
after they have been conditioned for 2-7 days in a 10%
CO2 atmosphere, the mean steady-state pHi of the
cell population was alkaline (Table I).
Rapidly growing HL60 cells that were constantly maintained in a 5%
CO2 incubator had average pHi of 7.27 ± 0.06. When these cells were conditioned in a 10% CO2 environment
and then placed back into medium pre-equilibrated with 5%
CO2 for 1 h, the mean pHi of the population
increased by about 0.12 pH units to 7.39 ± 0.07. If the
conditioned cells were resuspended in 5% CO2 medium for
2 h, pHi increased even further to 7.53 ± 0.09, similar to the steady state pHi of RV+ cells of
7.52 ± 0.03 (cf. Table I). Continued incubation past this time resulted in gradual return to base-line steady state pHi (not shown, but see Ref. 2). As described previously (2),
we interpret these changes in pHi to be the result of one or
more alkalinizing mechanisms that are up-regulated, while the cells are
cultured under an increased acid burden (i.e. 10%
CO2); when the acid burden is removed suddenly
(i.e. cells are returned to 5% CO2), these
alkalinizing mechanisms "overshoot" pHi until such time as
they are down-regulated as the cell readjusts to normal 5%
CO2. This down-regulation phase is typically not complete
for several hours (see Ref. 2 for detailed description).
Cl
When HL60 cells were pretreated with varying amounts of
H2DIDS before Cl Resistance of Alkaline HL60 Cells to CMC in Cl
HL60 cells with elevated pHi in Cl
HL60 cells were next pretreated with varying concentrations of
H2DIDS to test if the observed resistance to CMC was due to the Cl Elevation of pHi by CO2 Conditioning Also
Results in Resistance to CMC--
We also elevated the pHi of
control HL60 cells by conditioning them to a 10% CO2
atmosphere and then placing them back in a 5% CO2
environment in the presence of fresh pre-equilibrated RPMI medium (see
Ref. 2 for a discussion of CO2 conditioning). Conditioned
cells placed back in 5% CO2 for 2 h were then exposed to M195 and complement and found to be significantly resistant to
complement killing at a low doses of M195 antibody (Fig.
4B). The HL60 cells that were
grown continuously at 5% CO2 were effectively killed in a
dose-dependent manner as before. Importantly, the degree of
resistance to CMC after CO2 elevation of pHi was
time-dependent, as was the extent of alkalinization (see
"CO2 Conditioning Causes Alkalinization of pHi in
HL60 Cells"). When the 10% CO2-conditioned HL60 cells
were placed in 5% CO2-equilibrated medium for 1 h,
the cells were less resistant to CMC relative to HL60 cells that were
pulsed for 2 h in 5% CO2 (Fig. 4A). In cells pulsed for 2 h (Fig. 4B), observable resistance
extended to higher dosages of M195. Thus, more alkaline pHi
appears to result in higher levels of resistance to complement-mediated killing (compare Table I, Fig. 4). These data show that the resistance to CMC after alkalinization can be observed in cells alkalinized in
four different ways (chemotherapeutic drug selected with MDR expression; MDR gene transfection (4); Cl Kinetics of Modulation and Internalization of HL60 and
RV+ Cells Are Similar--
Since CMC is dependent first on
antibody binding and retention on the cell surface, especially in the
case of IgG antibodies, we wanted to ensure that the resistance of the
RV+ cells was not due to insufficient numbers of antibodies
binding or due to changes in modulation of the immune complex after
binding with consequent internalization. We found that
multidrug-resistant HL60/RV+ cells with elevated
pHi bind nearly twice as much 125I-HuM195 antibody
relative to parental HL60 cells, regardless of the length of time cells
are incubated with the antibody. These data confirm (in a more
quantitative fashion) our earlier conclusion that RV+ cells
display more cell surface antibody binding sites relative to parental
HL60 cells, extrapolated from indirect flow cytometry measurements (4).
The kinetics of cellular internalization of the antibody (revealed as
125I counts/min localized to the cell pellet) was similar
for both cell lines, although RV+ cells initially bound
greater levels of HuM195. Thus, importantly, any defect in initiation
of the terminal complement cascade (resulting in observed resistance to
CMC) cannot be due simply to reduced binding of relevant antibody to
the MDR cells; on the contrary, RV+ cells bound more immunoglobulin.
Kinetics of MAC Formation Is Delayed in Cells with Elevated
pHi--
Tschopp et al. (6) previously noted
that C9 polymerization required at least 10 min for completion under
conditions similar to those used in flow cytometry experiments. Thus,
to test whether elevation in pHi alone, without MDR
overexpression, affects binding of the MAC to the target cell membrane,
control HL60 cells were first CO2-conditioned and placed
back into 5% CO2 for 2 h to elevate pHi as
described above. Then MAC formation was measured by flow cytotometry
using an antibody that binds to a neoepitope found on C9 only after
binding to C8 or polymerizing to other C9 to form a mature MAC (Fig.
5). The alkaline HL60 cells exhibited a
delay in peak MAC binding as well as a marked reduction in MAC density
relative to control cells. The viability of the HL60 cells at 20 min
was 49%, while the "CO2-pulsed" HL60 cells had 84%
viability. The same experiment was repeated using P-GP-expressing RV+ cells (not shown). Over 30 min, mean peak channels for
the HL60 cells ranged from 3.3 to 7.3, while the RV+ cells
had mean peak channels of 2.6-3.9. The HL60 cells had a viability of
43% at the 20-min time point, while RV+ cells were 75%
viable.
Monitoring BCECF Leakage as a Measure of Functional MAC Pore
Formation--
Binding of a conformationally sensitive antibody is
informative but does not determine whether the MACs that are formed are functional. The electrophysiological assessment of MAC pore transport properties can be problematic. Therefore, we explored the possibility of following diffusion of a molecule with an effective radius near 2-3
nm (slightly below the upper limit pore diameter of a minimal
(C5b-C8)1(C9)9 full MAC; see Ref. 3) as an
indication of functional pore formation. We focused on the frequently
utilized fluorescence indicator BCECF. The acetoxy methyl ester
derivative of BCECF (BCECF-AM) is neutral and rapidly diffuses into
living cells, where it is cleaved by cytosolic esterases and obtains a
Upon the addition of antibody and complement proteins (serum
corresponding to
As described under "Experimental Procedures," the BCECF leakage
curve may be numerically fit to deduce key parameters. Differentiation converts the computed sigmoid to a gaussian, and the amplitude of the
gaussian (defined as A in Equation 1) represents the maximal rate of loss (Vmax) of the BCECF (slope of the
sigmoid at its midpoint; cf. Fig. 6A). The
characteristic width of the gaussian (
To test this method and our analysis, we performed multiple experiments
under identical conditions but varying the concentration of complement
(Fig. 6B). Titration with reduced amounts of complement (in
the presence of saturating antibody) significantly shifts the position
of the midpoint of maximal change in 439-nm BCECF fluorescence to the
right (i.e. increases the value of D, which reflects a decrease in the rate of pore formation). In contrast, only
mild effects are found for Vmax, suggesting (as
expected) that although the rate of formation of >2.5-nm diameter MACs
is dependent on the concentration of MAC subunits (Fig. 6B),
assuming the C9 component is not limiting, transport properties of the MACs that are formed (reflected by the value of
Vmax) are similar. Note that a significant
change in Vmax could (in theory) represent either a change in the number of MACs formed per cell or the intrinsic transport properties of the MAC (see below).
Following the method described above, we then compared 439-nm
fluorescence data for continuously mixed suspensions of HL60, multidrug
resistance HL60/RV+ cells, and alkalinized HL60 cells
preloaded to similar levels of BCECF and subsequently exposed to
similar levels of M31 antibody (Table II). For all cell lines, BCECF
leakage was dependent upon the concentration of complement added as
described in the legend to Fig. 6 (data not shown). However, D439 is
significantly larger for the RV+ cells, relative to the
control HL60 cells, at all values of complement tested (e.g.
Table II) although binding of antibody to RV+ is increased.
Thus, the rate of formation of BCECF-conducting ((C5b-C8)1(C9)9 (n Kinetics of MAC Formation Is Delayed in Cells with Elevated
pHi--
The increase in D439 values for the
RV+ and alkaline cells relative to HL60 suggests a longer
pore assembly time; however, changes in Elevated pHi Does Not Significantly Affect MAC Subunit
Stoichiometry--
Again, one likely cause of a possible change in MAC
pore size that has been previously described (29) would be a
significantly altered (C5b,C6,C7,C8)1(C9)n
stoichiometry. To assess subunit stoichiometry, dual isotope tracer
analysis of MAC components C7 and C9 was performed (see "Experimental
Procedures"). One C7 subunit is found per MAC, whereas C9 is
variable, with higher C9:C7 stoichiometry expected for pores of larger
diameter (29). Since there is only one C7 per MAC, quantifying labeled
C7 is an additional approach for estimating the number of MACs on the surface of each cell.
MAC molecular stoichiometry was determined following dual labeling of
complement components and purification of MACs in both HL60 cells in
HBSS and HL60 cells in Cl Resistance to CMC is modulated by P-GP overexpression (here and in
Ref. 4); therefore, a suitable molecular explanation for this
phenomenon must take into account the effects on agents acting
primarily on or in the cell plasma membrane (e.g. antibodies binding to the membrane surface and/or initial assembly of the MAC or
MAC pore function). Models envisioning that P-GP is an active efflux
pump for drugs (1, 3) or lipids (30) or is a "vacuum cleaner" (1)
do not easily account for the observed resistance to CMC.
Conversely, significant changes in pHi and
Vm have frequently been observed in MDR cells, are
due to P-GP overexpression (2, 9, 10), and more easily explain CMC resistance. Although two chemotherapeutic conditioned MDR cell lines
have been reported to have normal pHi (reviewed in Ref. 12), to
our knowledge, every MDR cell line that is MDR purely via
overexpression of P-GP (e.g. a "pure" phenotype not further complicated by exposure to chemotherapeutic drug) is alkaline, depolarized, or both (2, 12). These changes are sufficient to explain
the relatively low levels of chemotherapeutic drug resistance mediated
by P-GP overexpression alone (2). Moreover, these changes in
pHi (or Vm) might form the basis for a more
reasonable explanation for resistance to CMC, since significant
perturbations in these parameters could influence events that occur at
the membrane during initiation of the complement cascade
(e.g. assembly of the MAC). To test this idea, in the present study we devised two methods for stably elevating pHi in control HL60 cells over a period of time sufficient to measure CMC.
We then altered their pHi to alkaline values that approximate
those observed for the P-GP-overexpressing MDR derivative HL60
RV+ (4). These artificially alkaline HL60 were found to be
as CMC-resistant as were RV+ cells. Moreover, the degree of
alkalinization appeared to be related to the degree of resistance.
Resistance to CMC was also independent of the mechanism used to
alkalinize the cells (e.g. Cl Two exchangers present in most eukaryotic cell plasma membranes, anion
exchanger (Cl For example, in another recent study (35) an additional effect of P-GP
overexpression-induced elevations in pHi was found, namely
alterations in the kinetics of the apoptotic cascade for Chinese
hamster ovary fibroblasts. Rather dramatic decreases in pHi
occur very early on in apoptosis and may be casual or permissive for
subsequent events in the cascade (see Ref. 35 and references within).
Thus, pHi dysregulation caused by P-GP overexpression might
indeed be expected to perturb the kinetics of the apoptotic cascade, at
least for some cells. Therefore, the consequences of P-GP
overexpression should be viewed more broadly when discussing a variety
of clinically relevant resistance phenomena. The protection that
overexpression of the protein affords to different cell types under
different conditions appears to cover a much wider range of cytotoxic
situations than initially anticipated. From the clinical perspective,
this is of course extremely disappointing. Therapies designed to
circumvent P-GP function through modulating drug distribution (but
still dependent upon CMC or efficient induction of apoptosis) may not circumvent a MDR phenotype as efficiently as initially anticipated.
With regard to the molecular mechanism whereby elevated pHi
confers CMC resistance, we find that MDR HL60 cells and alkaline HL60
cells have similar binding kinetics for specific antibody at their
surface and that RV+ actually binds more
antibody; thus, resistance to CMC is not due to a reduced number of
antibody sites or reduced ability of antibody to bind (which would
provide fewer "anchors" for assembly of complement proteins and
subsequent cell lysis). However, flow cytometry utilizing a neo-C9
epitope suggested that the density of functional MACs on the cell
surface might be reduced. By analyzing the behavior of 439 BCECF
fluorescence traces, we are able to firmly conclude that the rate of
formation of complete MAC pores (C5b-C8)1(C9)n
(n = 9-18) is significantly reduced in the
RV+ and alkaline HL60 cells. Pore formation is slightly
faster in alkaline HL60 compared with RV+, suggesting that
additional changes in RV+ due to continued selection upon
chemotherapeutic drug may also contribute to defects in MAC assembly.
One possibility for these additional effects could be altered membrane
lipid composition, which appears to occur in some cells upon continued
exposure to chemotherapeutic drugs (30). Membrane changes caused by
chemotherapeutic drug selection could additionally affect MAC assembly
via perturbing electrostatic interactions that appear to be required
for insertion of C9 and other subunits (for example, changes in
sphingolipid/phosphatidylcholine ratios could alter surface charge,
which appears to play a role in regulating the protein conformational
changes that occur during C9 insertion (see below)).
Along with an apparent reduced rate of MAC formation, the BCECF leak
data also suggest a slower rate of cellular BCECF release (reduced
Vmax) once pores are formed for the MDR and
alkaline HL60 cells. Two explanations could conceivably account for
these results: 1) a decreased number of complete MAC pores are
assembled on the cell surface, or 2) the pore size of MACs for
RV+ and alkaline cells is smaller. By dual isotope labeling
experiments with purified complement components C7 and C9, we conclude
the subunit stoichiometry of MACs for HL60, RV+ and
alkaline HL60 is similar. However, more dramatically, quantitation of
C7 revealed that there was a clear, significant decrease in the number
of MACs assembled on the membrane of RV+ and alkaline
cells. This result supports that obtained by flow cytometric analysis
with a C9 antibody that binds to a conformationally sensitive
"functional" epitope expressed in the assembled MAC. Nonetheless, a
slight difference in C9 content is noted, and this could conceivably
contribute to the Vmax effect found for
RV+ and alkaline HL60. Although a one-C9 subunit-altered
stoichiometry could conceivably affect MAC pore diameter by as much as
9-10%, and although a smaller pore diameter would be consistent with a decreased Vmax, the large perturbations in
Vmax (70-75%) we observe are more easily
explained by the observed significant decrease in pore number.
Interestingly, Andrews and colleagues have observed that the
pore-forming protein from Trypanosoma cruzi, which was found to be immunologically related to human C9 forms "pores" (or
"complement channels") readily at lower pH but inefficiently at
higher pH (36). This observation is similar to those summarized in the present paper, wherein complete C9-containing pores are not formed as
efficiently at higher pHi.
Effects of extracellular pH on the activation or the pore
forming abilities of complement have also been observed, but these are
probably due to completely different mechanisms. Human C-reactive protein is known to activate the complement cascade upon reaction with
the complement component C1q (37). This activation is dramatically augmented when C-reactive protein is complexed with a suitable ligand,
such as phosphocholine-containing (38) and polycation-containing ligands (39). Miyazawa and Inoue (37) observed that in the absence of
phosphocholine-containing materials or polycations, the complement
cascade could be activated by C-reactive protein at mildly acidic
conditions (optimal at pH 6.3) in the presence of negatively charged
surfaces. Thus, both pHo and pHi are capable of
regulating CMC.
Esser et al. (40) have reported that the MAC is capable of
physically reorganizing the lipid bilayer during assembly and that the
forming MAC affects polar and nonpolar regions of the bilayers at
different stages of its assembly. They suggested that C5b-7 interacts
strongly with charged regions of the bilayer and may penetrate slightly
into the hydrophobic region, while C5b-8 and C5b-9 penetrate more
deeply into the membrane. Esser et al. (40) thus concluded
that during MAC formation the terminal components undergo dramatic
conformational change that allows specific binding sites for
phospholipids. Based on the results in this paper, we extend this
hypothesis to suggest that the conformational changes are highly
pHi-dependent or are dependent upon the magnitude of plasma membrane Finally, as interesting as these effects of pHi are, we note
that cells overexpressing human MDR 1 typically exhibit decreased
Vm as well as elevated pHi (Ref. 2 and
references therein). Possible additional effects (if any) of membrane
depolarization in conferring CMC resistance should also be investigated.
In sum, data in this paper and other recent studies (e.g.
Refs. 2 and 35) illustrate that models for P-GP function must accommodate a wider array of resistance phenomena than originally anticipated. We believe the altered partitioning model for P-GP (11,
12) is particularly helpful in this regard. It is a useful framework
for addressing both CMC resistance phenomena described within as well
as apoptotic cascade effects (35) and the thermodynamic and kinetic
riddles associated with altered drug partitioning in MDR cells
(reviewed in Ref. 12). Further inspection of the model's implications
should continue to resolve clinical questions.
substitution), are
tested for CMC using two different antibody-antigen systems (human IgG
and murine IgM; protein and carbohydrate) and two complement sources
(rabbit and human). Elevation of pHi by either of these methods
or by expression of P-GP confers resistance to CMC. Resistance is not
observed when the alkalinization mediated by reverse
Cl
/HCO3
exchange upon
Cl
substitution is blocked by treatment with
dihydro-4,4'-diisothiocyanostilbene-2,2'-disulfonate. Continuous
photometric monitoring of
2',7'-bis(carboxyethyl)-5,6-carboxyfluorescein (BCECF), to assess
changes in pHi or efflux of the probe through MAC pores, in
single cells or cell populations, respectively, verifies changes in
pHi upon CO2 conditioning and Cl
substitution and release of BCECF upon formation of MAC pores. Antibody
binding and internalization kinetics are similar in both the parental
and resistant cell lines as measured by radioimmunoassay, but flow
cytometric data showed that net complement deposition in the cell
membrane is both delayed and reduced in magnitude in the MDR cells and
in the cells with increased pHi. This interpretation is
supported by comparison of BCECF release data for the different cells.
Dual isotopic labeling of key complement components shows no
significant change in molecular stoichiometry of the MACs formed at
different pHi. The results are relevant to understanding
clinical implications of MDR, the physiology of P-GP, and the
biochemistry of the complement cascade and further suggest that the
"drug pump" model of P-GP action cannot account for all of its effects.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
directly or indirectly. Cytosolic
Cl
concentration and Cl
plasma membrane
permeability are generally important in maintaining pHi and
Vm for eukaryotic cells. Altered Cl
permeability and altered pHi in MDR cells may indeed be
connected in some fashion, since recent Cl
substitution
experiments revealed unusual and extensive pHi changes in human
MDR 1 transfectants (16). Overexpression of P-GP inhibits
Cl
and
HCO3
-dependent pHi
homeostasis, and this is apparently due to specific inhibition of (or
competition against) Na+-independent
Cl
/HCO3
exchange (anion
exchange (10, 16, 17)). HL60 cells are known to possess
Cl
/HCO3
exchange
activity (18), so P-GP expression in these cells would be expected to
perturb pHi.
/gluconate
exchange and CO2 conditioning) were used to alkalinize control HL60 cells not expressing P-GP. Intracellular alkalization of
HL60 cells conferred CMC resistance similar to that measured for the
RV+ cells expressing P-GP. Resistance did not develop when
intracellular alkalinization upon Cl
/gluconate exchange
was prevented via inhibition of the
Cl
/HCO3
exchanger due to
the addition of dihydro-4,4'-diisothiocyanostilbene-2,2'-disulfonate (H2DIDS). Furthermore, the pHi changes did not
appear to change antibody binding and modulation kinetics but did
appear to have effects on complement membrane attack complex (MAC) deposition.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
substitution as described (10, 16, 24). After a 10-min perfusion,
normal HBSS perfusate was rapidly replaced with Cl
-free
HBSS (gluconate isotonically replacing Cl
). Control
experiments (not shown) showed that excess glutamate tended to inhibit
the complement cascade (see Ref. 26), so gluconate was preferred in
isotonic Cl
substitution. Long term Cl
substitution experiments performed essentially as described (16) verified that HL60 cells remained at an alkaline pHi under
these perfusion conditions during the duration of a typical complement
cytotoxicity assay (about 30 min; see "Results").
substitution conditions,
H2DIDS, a well known anion exchange inhibitor, was added to
the cells at various concentrations (250 µM to 1 mM) after loading with BCECF. The cells were incubated with
the H2DIDS for 20 min at 37 °C, and isotonic
Cl
/gluconate exchange was performed as above in the
presence of CO2/HCO3
to
examine any inhibition of
Cl
/HCO3
exchange (see
"Results").
-free
HBSS--
Complement-mediated cytotoxicity assays were conducted
essentially as described by Weisburg et al. (4) except that
cells were washed to remove all medium and resuspended in HBSS
pre-equilibrated to 37 °C and pH 7.30 (by perfusion with 5%
CO2). The cells were allowed to equilibrate in HBSS
containing 5% fetal calf serum for 10 min and then plated for the
cytotoxicity measurements (4). For the Cl
-free HBSS
experiments, the cells were pretreated with HBSS as above, washed with
Cl-free HBSS to remove Cl
, and then resuspended in
Cl
-free HBSS at 37 °C, pH 7.30, with 5% fetal calf
serum. Antibody and complement dilutions were made in either HBSS or
Cl
-free HBSS as appropriate.
/HCO3
exchanger function was involved in any resistance to complement when
the cells were suspended in Cl
-free HBSS, cells were
treated with H2DIDS prior to the CMC assay. HL60 cells were
washed free of medium, resuspended in HBSS plus H2DIDS
(concentrations ranged from 250 µM to 1 mM),
and then incubated for 20 min at 37 °C in an atmosphere of 5%
CO2. The cells were washed with Cl
-free HBSS
and then plated for cytotoxicity with an amount of antibody and
complement known from previous experiments to yield approximately 50%
killing (see "Results").
-free Conditions--
We modified the protocol
previously published by Ware and colleagues (28). 5.0 × 108 HL60 cells were washed three times with HBSS, pH 7.30, to remove all growth media. Next, the cells were resuspended in HBSS
plus 10% fetal bovine serum and incubated for 10 min at 37 °C. For Cl
-free conditions, the HL60 cells were then washed two
times with Cl
-free HBSS (equimolar gluconate substituted
for Cl
) plus 10% fetal bovine serum and incubated at
37 °C for 5 min to allow the pHi of the cells to stabilize.
After this incubation period, cells were collected by centrifugation
and resuspended in a 1:13 dilution of M31 monoclonal antibody and a 1:3
dilution of human serum as a complement source. A tracer amount of
131I-C7 (9.50 × 108 cpm, specific
activity = 2.80 × 104 cpm/ng) and
125I-C9 (8.50 × 108 cpm, specific
activity = 3.20 × 104 cpm/ng) was also added.
The cells, antibody, complement, and radiolabeled complement components
were incubated at 37 °C for 35 min. After this incubation, cell
viability was determined by trypan blue exclusion. Relaxation buffer
(2.5 M KCl, 1 M MgCl2, 1 M NaCl, 0.5 mM ATP, 0.5 M Tris-HCl,
pH 7.6) plus 2 mM phenylmethylsulfonyl fluoride and
aprotinin to a final concentration of 100,000 units/ml was added to the
cells. The flask of cells was then placed into a nitrogen bomb and
pressurized to 350 p.s.i. The cells were lysed by nitrogen
cavitation, and unlysed cells were collected by centrifugation at
750 × g. The cell membrane fraction was resuspended in
BEP buffer (5 mM sodium borate, 10 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, pH 8.8) and washed three
times. Protein concentrations were determined by the Bio-Rad protein
assay. The washed HL60 cell membranes were extracted with 1%
deoxycholate in BEP buffer (40:1 detergent:protein ratio (w/v) for 30 min at 25 °C with mixing every 5 min). Cell membranes were collected
by centrifugation at 27,000 × g for 30 min; the
clarified supernatant fluid containing the MACs was concentrated to
less than 2 ml by centrifugation in a Centriprep 50 spin column
(Amicon, Inc., Beverly, MA) and subjected to gel filtration
chromatography (Biogel A-15M column size 2 × 60 cm) in the
presence of 50 mM Tris acetate, pH 8.8, and 1%
deoxycholate. Fractions were collected in 2-ml volumes. Protein was
monitored by absorbance at 280 nm, and radioactivity was measured for
aliquots of the fractions by liquid scintillation spectrometry (Packard).
-free HBSS as appropriate and resuspended in 0.5 ml of
the same buffer. The cells were then placed in a rapidly stirred
cuvette containing 2 ml of the appropriate buffer. The buffers were
continuously purged with 5% CO2 to maintain extracellular
pH, which was monitored with a microelectrode. The cells were
equilibrated with the HBSS or Cl
-free HBSS for 10 min,
and the baseline 439/490-nm BCECF excitation ratio was monitored with
an alpha-scan fluorometer (Photon Technology Inc., New Brunswick, NJ).
After a flat base line was verified, 100 µl of dialyzed M31 antibody
(approximately 2.5 µg total) was added. The cells were incubated with
the antibody for 100 s, during which time no appreciable change in
base-line fluorescence was noted (see "Results"). Finally, variable
rabbit complement was added (see "Results"). BCECF leakage through
large complement pores was determined by the increase in fluorescence
excitation at 439 nm as the BCECF exited the cells (due to an increase
in quantum yield for solution-based BCECF versus
intracellular BCECF; see "Results"). Changes in 439-nm fluorescence
are not expected upon any possible movement of H+ during the
experiment, since 439 nm is isosbestic with regard to changes in pH.
This was confirmed by titration of BCECF in the cuvette (not shown).
Parallel experiments wherein cells and supernatant were separated and
fluorescence in each fraction was quantified individually (not shown,
but see Ref. 9) verified that the change in 439-nm BCECF fluorescence was indeed due to exit of the probe from the cell. As an additional control experiment, 200 µl of heat-inactivated rabbit complement was
added to the cells plus M31 antibody (see "Results") in order to
test whether any change in probe signal was caused by the addition of
reagents but in the absence of functional pores.
which is the integral of a gaussian (i.e. a sigmoid),
and where F(t) is fluorescence intensity at 439 nm excitation (or the 439/490-nm excitation ratio, see "Results")
as a function of time (t); and A, C,
(Eq. 1)
, and tm are variables. The expression under the
integral has the general form of a bell-shaped (Gaussian) curve. After
iterative least-squares best fit to the raw data (convergence satisfied
in <100 iterations; p < 0.05), the expression perfectly fit the raw data (see "Results"). In this expression, C corresponds to the intensity of fluorescence before
complement is added, A is the maximal rate of change in
F(t) (e.g.
Vmax, which is observed at time
tm), and
defines the characteristic time of
transition of F(t) to a new steady state value.
We also define the "delay time" (D) in the equation,
where t0 is the time at which complement
is added to the cell suspension; i.e. D is the
time between the addition of complement and the maximal rate of change
in F(t). These simple parameters are useful for
quantifying the rate of formation of pores (i.e. the
"delay time", D) as well as the cellular rate of
transport (maximal rate of change, A, also referred to as
Vmax), which is proportional to the number of
pores formed and/or the intrinsic transport properties of those pores
that are formed.
(Eq. 2)
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
Steady state pHi for HL60 cells
-free
medium for 5 min. Measurements were performed on at least 15 individual
cells, in triplicate (45-60 cell total) via single cell photometry as
described under "Experimental Procedures" using three separate
coverslips under constant perfusion (see "Experimental Procedures";
also described in detail in Ref. 2).
/Gluconate Exchange Increases pHi of
HL60 Cells--
Upon exposure to Cl
-free HBSS, HL60
cells with an initial pHi near 7.25 became significantly
alkaline, with pHi increasing to near 7.65 (Fig.
1A). This alkalinization is
dependent upon the presence of HCO3
(not shown) and is inhibited by H2DIDS (see Fig.
1B). Previously (18), Restrepo and colleagues demonstrated
the presence of a Cl
/HCO3
exchanger in
HL60 cells. Thus, it is likely that alkalinization is mediated by
"reverse" Cl
/HCO3
exchange instigated by the outward directed Cl
flux
caused by isotonic Cl
substitution (see Ref. 16).
Interestingly, HL60 remained stably alkaline for at least 30 min in
Cl
-free HBSS. This suggests that HL60 cells lack a
compensatory Cl
-independent mechanism capable of
restoring normal pHi under these conditions (e.g. a
Na+/HCO3
cotransporter).
Stability of alkalinization under these conditions is important, since
assaying complement-mediated cytotoxicity in this system requires a
30-min incubation period with complement (see below).
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Fig. 1.
Intracellular pH for single HL60 cells under
constant perfusion. Cells were attached to glass via the Becton
Dickenson "Cell Tak" method (see "Experimental
Procedures"). After perfusion with HBSS for 5 min, pHi was
monitored, and isotonic Cl /gluconate exchange was
performed (first arrow at 250 s).
A, this protocol elevates pHi due to the influx of
HCO3
coupled to Cl
efflux via the anion exchanger present in these cells (described in
Ref. 18). B, alkalinization in H2DIDS0-treated
HL60 is markedly reduced under similar conditions, due to inhibition of
the Cl
/HCO3
exchanger by
H2DIDS. Pretreatment with the stilbene also appears to
lower resting pHi slightly. After verifying that alkalinization
is reasonably stable for approximately 30 min under each condition,
cells were titrated to pHi 6.80, 7.20, and 7.60 (second, third, and fourth
arrows, respectively) to calibrate internal BCECF response
for the same cell (see Refs. 24 and 25). Data shown in these
panels are representative of many experiments
(n > 12 for each cell type) that were averaged for
different cells to produce the data shown in Table I.
/gluconate exchange, the
cells did not become significantly alkaline (Fig. 1B). The
initial pHi of the H2DIDS-treated HL60 cells
(pHi 7.07 ± 0.09) is lower than untreated HL60 cells,
suggesting that a stilbene-inhibitable transport process is required to
maintain normal steady state pHi. In any case, inhibition of
substantial alkalinization by H2DIDS (Fig. 1, compare
A and B) further supports the notion that
alkalinization upon Cl
substitution is largely due to
reverse Cl
/HCO3
exchange
mediated by the Cl
/HCO3
exchanger present in these cells (18).
-free
HBSS--
Monoclonal antibody M195 reacts with an antigen typically
expressed at a density of about 10,000 sites/cell (GP 67). HL60 cells
with elevated pHi due to isotonic Cl
substitution
(Fig. 2A, gray
bars) were resistant to M195 IgG1-mediated CMC
as compared with HL60 cells suspended in normal HBSS (Fig. 2A, open bars). The degree of
resistance for the alkaline cells is similar to the degree of
resistance seen previously for the MDR variant RV+ cells
(Ref. 4 and Fig. 2A, solid bars),
which also exhibit an elevated pHi relative to control HL60
cells (Table I). The differences in killing for the alkaline
versus control cells are most prominent at lower
concentrations of complement and antibody. This suggests that
resistance can be overcome with increasing antibody.
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Fig. 2.
Complement-mediated cytotoxicity in HL60
(open bars), HL60 RV+
(solid bars), and HL60 in
Cl -free medium (gray
bars) using monoclonal antibody M195 and rabbit
complement (A) or monoclonal M31 and human complement
(B). Data shown are the averages from a single
experiment (each sample done in duplicate) and are representative of
many experiments. Controls (with antibody and without complement, or
with complement and without antibody; shown at right) verify
killing is via the CMC pathway.
-free HBSS were
also more resistant to a wide range of effective concentrations of an
IgM (M31) that recognizes the carbohydrate epitope Lewis X (CD15), in
the presence of human complement (Fig. 2B). Again, the
efficiency of M31-mediated CMC for HL60 in Cl
-free HBSS
(Fig. 2B, gray bars) closely resembled
that observed for the MDR RV+ cells (Fig. 2B,
solid bars).
/HCO3
exchange-dependent increase in pHi and not merely
to the absence of Cl
. In these experiments, the
concentration of M195 or M31 that yielded the approximate
LD50 (determined in the previous experiment, shown in Fig.
2) was used along with either rabbit complement or human serum,
respectively. On average, 44% HL60 cells in HBSS were killed by CMC
(Fig. 3A, open
bars), whereas only 8% of the HL60 cells in
Cl
-free HBSS (Fig. 3A, solid
bars) were killed by CMC at this dose of antibody. HL60
cells pretreated with 1 mM H2DIDS before being resuspended in Cl
-free HBSS (Fig. 3A,
stippled bars) showed a sensitivity similar to
that of HL60 in HBSS. Control experiments in which HL60 cells in
Cl
-free HBSS and treated with only 1 mM
H2DIDS, using medium lacking either complement or antibody
(heavy stippled and striped
bars, respectively; Fig. 3A), were not killed
efficiently showed that the increased cytotoxicity was not due to
treatment with H2DIDS in and of itself (Fig. 3A,
compare third, fourth, and fifth
bars from the left). Similar relative results
were also obtained at lower H2DIDS concentrations, where
the effects of the stilbene are predicted to be even more specific to
Cl
/HCO3
exchanger
proteins (not shown). A similar reversal of resistance to CMC was seen
for H2DIDS-treated HL60 in Cl
-free HBSS that
were exposed to M31 IgM antibody and human complement (Fig.
3B).
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Fig. 3.
Complement-mediated cytotoxicity on HL60
cells before (open bars) or after isotonic
Cl substitution to elevate pHi (solid
bars) or after isotonic Cl
substitution
after pretreatment with H2DIDS (gray
bars). M195 (A) or M31 (B)
was the antibody used to initiate CMC. Cells exposed to
Cl
-free conditions but not H2DIDS
(solid bars) are alkaline (Table I) and resistant
to CMC, whereas those also pretreated with H2DIDS
(lightly stippled bars) are not. Since
treatment with H2DIDS in the absence of an initiated CMC
pathway (e.g. in either the absence of antibody or
complement; heavy stippled and striped
bar; on far right) does not result in
effective kill, reversal of resistance by H2DIDS (a potent
inhibitor of Cl
/HCO3
exchange in HL60 cells) indicates that the resistance to CMC upon
Cl
substitution (solid bar) is due
to alkalinization caused by the influx of
HCO3
under these conditions (see Fig.
1A).
substitution;
and CO2 pulse) using either human or mouse immunoglobulins of IgG and IgM isotypes and utilizing either rabbit or human
complement.
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Fig. 4.
Comparison of CMC for HL60 grown continuously
at 5% CO2 (open bars),
RV+ grown at 5% CO2 (solid
bars), and HL60 "pulse-elevated" in
pHi by 10% CO2 conditioning followed by placement
at 5% CO2 (stippled bars) for
either 1 (A) or 2 h (B).
Relative alkalinization by this method is dependent upon the time at
5% CO2 (see also Ref. 2) as is the relative resistance to
CMC.
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Fig. 5.
Flow cytometric analysis of MAC pore
formation (kinetics) using an antibody that recognizes the neoepitope
on C9 during mature MAC formation but not the free C9 component.
Normal HL60 cells had an increasing mean peak channel (functional MACs)
over time. The CO2-pulsed HL60 cells showed no increase in
mature MAC binding during the same time period. Mean peak channel
correlates with the number of C9 neoepitopes per cell.
4 charge. Passive diffusion of converted BCECF from the cytosol of
cells with no large membrane pores is very slow (at most, 5%/h under
constant perfusion conditions; see Ref. 2), yet importantly, we do
observe significant differences in the effective quantum yield of
fluorescence for intracellular versus extracellular BCECF (see Fig. 6A). Moreover, since
the fluorescence excitation spectrum is complex, it is reasonable to
anticipate that monitoring multiple peaks representing different
transitions that respond differently to changes in solvent polarity is
feasible. Thus, similar to other studies using a "continuous
monitoring of fluorescence" approach (9) but using constantly mixed
suspensions of BCECF-preloaded cells (see "Experimental
Procedures"), we followed the loss of intracellularly trapped BCECF
via a change in the quantum efficiency of 439-nm excitation (Fig.
6A) as a measure of MAC pore formation.
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Fig. 6.
A, 439-nm BCECF fluorescence data for
continuously mixed suspensions of HL60 cells preloaded with BCECF-AM to
which normal complement proteins (upper curve) or
heat-inactivated complement (lower line) was
added 100 s after the addition of M31 antibody (antibody added at
the arrow). In each case, 1.0 × 106 cells
preloaded with BCECF-AM for 30 min were used (see "Experimental
Procedures"). The sigmoidal data (top curve) is
easily fit to Equation 1 as described under "Experimental
Procedures." Data shown are representative of dozens of experiments.
In general, at this dose of complement, BCECF leak (revealed as the
increase in excitation efficiency) began within 12-13 min after the
addition of complement proteins. B, representative 439-nm
fluorescence data obtained for aliquots of control HL60 cells treated
under identical conditions but exposed to variable levels of complement
(from left, serum complement that includes 12, 6, or 3 µg/ml C9). Upon diluting complement 2-fold, the time before the
appearance of 439-nm changes (illustrating formation of full MACs and
loss of intracellular BCECF) increases substantially. A plot of
computed "delay time" (D) for these curves and others
versus complement concentration yields a quadratic
relationship (not shown, but see data in Table II) that plateaus near
10 min.
15 µg/ml C5, C6, C7, and C8 and approximately 12 µg/ml C9 final
concentration)4 to cells,
loss of intracellularly trapped BCECF (seen as an increase in 439-nm
excitation efficiency for the continuously mixed cell suspension)
begins within 12-13 min (Fig. 6A; see also Table
II). This time is similar to the time at
which the first indications of altered cell viability via trypan blue
exclusion are evident (data not shown, but see Ref. 4). By physically
separating cells from the medium at various times after the addition of
complement and accounting for scatter from the cell suspension (not
shown) we are able to conclude that a 2.0-2.5-fold change in 439-nm
excitation quantum efficiency accompanies the release of trapped BCECF
from these cells and that virtually all BCECF is released within 30 min
at this concentration of complement and antibody (see Ref. 9 for
similar calculations). No change in 439-nm BCECF response is seen for
the continuously mixed suspension when an equivalent amount of
heat-inactivated complement is added (Fig. 6A,
bottom trace) or when nigericin is added to
intentionally change pHi (not shown), further supporting the
conclusion that these data illustrate formation of large functional
(>2.5-nm diameter) pore MACs that accommodates the diffusion of
BCECF.
Summary of curve fitting analysis for 439-nm BCECF data
2 and Vmax is relative to the control
HL60 at 12 µg/ml C9. Notably, D439 values are significantly delayed
for RV+ relative to HL60, suggesting a significant increase in
the rate of formation of BCECF-conducting MAC pores. Similarly,
alkaline HL60 also exhibits increased D values (see "Results").
2) is the time it
takes for all BCECF to be released after pores are formed (time of
transition between the two steady states), and the difference between
t0 (time of the addition of complement, denoted
by the arrow in Fig. 6A) and
tm (time at which Vmax is
computed, i.e. the midpoint of the sigmoid) is defined as
the "delay time" (D) or mean time required for fully
BCECF-competent channels to form "complete MACs" (Table II).
9))
functional MAC pores is dramatically reduced for the MDR derivatives.
Similarly, Vmax was slower for RV+
(approximately 3-fold), suggesting either a reduced number of complete
MACs or a change in their intrinsic transport properties (this
distinction will be discussed below). These defects probably explain
resistance to CMC. Similar alterations in the formation of functional
MAC pores were also found for HL60 cells with elevated pHi
induced by the Cl
substitution method (Table II). Similar
to the case for RV+, D439 values are larger at a given
concentration of complement, and Vmax values are
lower, suggesting a reduced rate of MAC formation and either a reduced
number of pores or altered MAC transport properties (see below).
2 and
Vmax have multiple explanations. Two
possibilities are as follows: 1) there are fewer functional MACs on the
surface of the cells with the higher pHi, or 2) the relative
permeability of MAC pores that are formed is decreased. A likely
molecular explanation for possibility 2 would be altered
(C5b,C6,C7,C8)1(C9)n stoichiometry, which would
change pore diameter. Data in Fig. 5 that quantify binding of an
antibody that binds to a conformationally dependent epitope on C9
suggested slower deposition of functional MAC complexes in cells with
elevated pHi. To further distinguish between these
possibilities, we analyzed the formation of functional (polymerized C9)
MACs in live cells and purified MACs after dual isotope labeling of
purified MAC components C7 and C9.
-free HBSS (Fig.
7). The isolation of the principal
fraction of intact MACs by size exclusion chromatography was resolved
at similar elution times with both cell populations (Fig. 7,
A and B). A minimal amount was eluted as a broad
peak (probably representing partially dissociated MAC complexes lacking
one or more subunits) resolved at later elution times. The majority of
125I-C9 and 131I-C7 radioactivity (Fig. 7,
compare solid line to dotted) eluted with the major protein peak upon MAC size exclusion chromatography. To
ensure that the principal peak of radioactivity represented labeled
complement components incorporated into the MAC and not free individual
complement components, labeled individual components were also added to
the column with human serum but without antibody to fix the complement.
Free C7 and C9 were both resolved with much longer elution times
relative to the radioactivity peak shown in Fig. 7, indicating that
this peak represents C7 and C9 complexed into the MAC. Quantitation
from these experiments performed with HL60 cells in HBSS showed that
the MACs exhibit an average C7:C9 ratio of 1:9.7 and that there were
approximately 25,000 MACs/cell membrane (55% of cells lysed). For HL60
cells resuspended in Cl
-free HBSS to alkalinize
pHi, the average C7:C9 ratio was 1:8.7, and there were only
approximately 10,400 MACs/cell membrane (only 25% of cells lysed.
Hence, apparent pore size is only slightly affected for HL60 cells with
an elevated pHi, but a much more significant reduction in MAC
density is clearly apparent.
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Fig. 7.
Size exclusion chromatography of intact MACs
purified from HL60 plasma membranes via the procedure of Ware et
al. (28). A, the 125I-C9 elution
profile on HL60 cells in HBSS; B, the 131I-C7
elution profile on HL60 cells in HBSS; C, the
125I-C9 elution profile on HL60 cells in
Cl -free HBSS; D, the 131I-C7
elution profile on HL60 cells in Cl
-free HBSS for
representative columns. In each case, the open
squares summarize data for membranes isolated under
conditions where MACs are expected to be present (see "Experimental
Procedures" and "Results"), and the open
diamonds represent control data for uncomplexed (free)
complement proteins where MACs are not found. Free C9 and C7 (via
either 125I or 131I data) elute near fractions
45-47, whereas complexed C9 and C7 elute in fractions 28-32. By
knowing specific activity and ratioing 125I
versus 131I, (C5b,C6,C7,C8):(C9) stoichiometry
can be computed.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
substitution
versus CO2 pulse versus P-GP
overexpression (4)). To firmly test that data from the Cl
substitution experiments indicated that elevation of pHi and
not simply loss of Cl
caused resistance to CMC, the
effect of Cl
removal in the presence of the stilbene
H2DIDS (to inhibit intracellular alkalinization via reverse
Cl
/HCO3
exchange) was
examined, and the alternative less invasive CO2 conditioning technique was also used. Similar resistance to CMC was
observed when pHi was elevated via CO2 pulse,
regardless of the specific antibody isotype, antigen type, or species
of complement. Thus, elevated pHi in MDR cells (caused by P-GP
overexpression) is the most likely explanation for the phenomenon of
resistance to CMC.
/HCO3
exchanger) and Na+/H+ exchanger, are vital to pHi
homeostasis. In cells that express P-GP, most of the available data
suggest that observed alkaline pHi is probably not due to
disruption of Na+/H+ exchanger but to dysregulation of
Cl
-dependent pHi homeostasis (24).
Another hypothesis for the function of P-GP is based on observations of
altered ATP transport in MDR cells (31). In this model, ATP transported by P-GP could activate a purinergic receptor/G protein signal transduction pathway that could then regulate ion channels (32). External ATP can increase [Na]i and induce biphasic
pHi changes characterized by transient acidification followed
by significant and sustained alkalinization (33, 34). However,
ATP-induced alkalinization of pHi is thought to be due to
activation of the amiloride-sensitive Na+/H+ exchanger,
which, as mentioned, is probably not responsible for P-GP-mediated
increases in pHi (16, 24). Regardless, via whatever mechanism,
elevated pHi caused by overexpression of P-GP may lead to more
and different phenotypic alterations in MDR cells than was previously
believed to be the case. Resistance to CMC is one example of this.
Variability in these phenotypic characteristics for different cell
types might help to explain the wide heterogeneity in MDR phenotypes
observed to date.
pH.
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ACKNOWLEDGEMENTS |
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We thank Dr. J. R. Bertino
(Sloan-Kettering Institute, New York) for the use of the -scan
fluorometer and Dr. M. M. Hoffman (Allegheny University of the
Health Sciences, Philadelphia, PA) for help with tissue culture.
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FOOTNOTES |
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* This work was supported by National Institutes of Health Grants CA55349 and GM54516.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.
¶ To whom correspondence should be addressed. Memorial Sloan-Kettering Cancer Center, 1275 York Ave., New York, NY 10021. Tel.: 212-639-8603.
2 D. A. Scheinberg and M. Tanimoto, unpublished results.
3 S. Dzekunov, J. Weisburg, D. A. Scheinberg, and P. D. Roepe, manuscript in preparation.
4 The approximate concentrations of complement proteins are upper limit estimates (41). For (C5b,C6,C7,C8)1(C9)n (n = 9-18) MAC stoichiometry, the C9 subunit is by far the limiting component.
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ABBREVIATIONS |
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The abbreviations used are: MDR, multidrug resistance; pHi, intracellular pH; CMC, complement-mediated cytotoxicity; P-GP, P-glycoprotein; MAC, membrane attack complex; Vm, membrane potential; BCECF-AM, 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein; AM, acetoxymethyl ester; HBSS, Hanks' balanced salt solution; PBS, phosphate-buffered saline; H2DIDS, dihydro-4,4'-diisothiocyanostilbene- 2,2'-disulfonate.
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REFERENCES |
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