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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 92 No. 12 (December 15), 1998:
pp. 4844-4855
Two Distinct Pathways Mediate the Formation of Intermediate Density
Cells and Hyperdense Cells From Normal Density Sickle Red Blood
Cells
By
Robert S. Schwartz,
Sylvia Musto,
Mary E. Fabry, and
Ronald L. Nagel
From The Albert Einstein College of Medicine and Montefiore Medical
Center, Bronx Comprehensive Sickle Cell Center and Division of
Hematology, Bronx, NY.
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ABSTRACT |
In sickle cell anemia (SS), some red blood cells dehydrate, forming
a hyperdense (HD) cell fraction (>1.114 g/mL; mean corpuscular hemoglobin concentration [MCHC], >46 g/dL) that contains many irreversibly sickled cells (ISCs), whereas other SS red blood cells
dehydrate to an intermediate density (ID; 1.090 to 1.114 g/mL; MCHC, 36 to 46 g/dL). This study asks if the potassium-chloride cotransporter
(K:Cl) and the calcium-dependent potassium channel [K(Ca2+)] are participants in the formation of one or
both types of dense SS red blood cells. We induced sickling by exposing
normal density (ND; 1.080 to 1.090 g/mL; MCHC, 32 to 36 g/dL) SS
discocytes to repetitive oxygenation-deoxygenation (O-D) cycles in
vitro. At physiologic Na+, K+, and
Cl , and 0.5 to 2 mmol/L Ca2+, the
appearance of dense cells was time- and pH-dependent. O-D cycling at pH
7.4 in 5% CO2-equilibrated buffer generated only ID cells,
whereas O-D cycling at pH 6.8 in 5% CO2-equilibrated buffer generated both ID and HD cells, the latter taking more than 8 hours to form. At 22 hours, 35% ± 17% of the parent ND cells were
recovered in the ID fraction and 18% ± 11% in the HD fraction.
Continuous deoxygenation (N2/5% CO2) at pH 6.8 generated both ID and HD cells, but many of these cells had multiple
projections, clearly different from the morphology of endogenous dense
cells and ISCs. Continuous oxygenation (air/5% CO2) at pH
6.8 resulted in less than 10% dense cell (ID + HD) formation. ATP
depletion substantially increased HD cell formation and moderately
decreased ID cell formation. HD cells formed after 22 hours of O-D
cycling at pH 6.8 contained fewer F cells than did ID cells, suggesting that HD cell formation is particularly dependent on HbS polymerization. EGTA chelation of buffer Ca2+ inhibited HD but not ID
cell formation, and increasing buffer Ca2+ from 0.5 to 2 mmol/L promoted HD but not ID cell formation in some SS patients.
Substitution of nitrate for Cl inhibited ID cell
formation, as did inhibitors of the K:Cl cotransporter, okadaic acid,
and [(dihydroindenyl) oxy]alkanoic acid (DIOA). Conversely, inhibitors of K(Ca2+), charybdotoxin and
clotrimazole, inhibited HD cell formation. The combined use of
K(Ca2+) and K:Cl inhibitors nearly eliminated dense cell
(ID + HD cell) formation. In summary, dense cells formed by O-D
cycling for 22 hours at pH 7.4 cycling are predominately the ID type,
whereas dense cells formed by O-D cycling for 22 hours at pH 6.8 are
both the ID and HD type, with the latter low in HbF, suggesting that HD
cell formation has a greater dependency on HbS polymerization. A
combination of K:Cl cotransport and the K(Ca2+)
activities account for the majority of dense cells formed, and these
pathways can be driven independently. We propose a model in which
reversible sickling-induced K+ loss by K:Cl primarily
generates ID cells and K+ loss by the
K(Ca2+) channel primarily generates HD cells. These
results imply that both pathways must be inhibited to completely
prevent dense SS cell formation and have potential therapeutic
implications.
© 1998 by The American Society of Hematology.
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INTRODUCTION |
THE ORIGIN OF dehydrated sickle cells,
either dense discocytes or irreversibly sickled cells (ISCs), found in
sickle cell disease (SS) blood has been a challenge since first
observed by Herrick nearly 90 years ago.1 Dehydration of SS
red blood cells has pathophysiological consequences, including
increased red blood cell rigidity2 and reduced life
span.3 Moreover, dehydration of SS cells implies an
increased mean corpuscular hemoglobin concentration (MCHC) that
decreases the delay time for deoxygenation-induced polymerization of
hemoglobin S (HbS) by a factor that is proportional to the 30th power
of the Hb concentration.4 Dense SS cells are also
rheologically incompetent5,6 and contribute directly to
vaso-occlusion.7
SS cells are heterogeneous with respect to degree of dehydration, age,
HbF levels, and ion transport properties.8,9 Some intermediate density (ID) cells (similar to the SS3 fraction previously defined by us as all red blood cells with MCHC of 37 to 42 g/dL) are
dense discocytes, whereas a hyperdense (HD) fraction (similar to the
SS4 fraction previously defined by us as all red blood cells with MCHC
>42 g/dL) is the class of SS cells that contains the majority of ISCs
as well as very dense discocytes.5
There are three operational ways to define dense cells in sickle cell
anemia. The first is to define them as all red blood cells more dense
than most normal (AA) red blood cells, which results in a density cut
between 1.091 and 1.096 g/mL. The second is to define a more dehydrated
group of SS cells that are rich in ISCs and depleted of HbF that are
the result of a density cut between 1.103 and 1.118 g/mL. In this
report, we have opted for a third approach. We have defined two classes
of dense SS cells: ID cells (1.090 to 1.114 g/mL) and HD cells
(>1.114 g/mL, which includes the ISC-rich cell fraction).
This approach is not new, because Clark et al10 proposed
similar criteria for cutting discontinuous Stractan gradients; red
blood cells of middle density began at 1.096 g/mL and hyperdense cells
began at 1.115 g/mL. Other workers have also defined two classes of
dense SS cells with cuts at 1.097 and 1.104 g/mL,11 1.087 and 1.118 g/mL,12 and 1.098 and 1.112 g/mL.13
Densities exceeding 1.105,14 1.110,15 and 1.118 g/mL16 have been used to define a single class of dense SS
cells. Others have made a broader definition of dense SS cells that
included all cells greater than 1.092 g/mL17 or greater
than 1.096 g/mL.18
In our previous work with continuous Percoll-Larex (Stractan)
gradients,19,20 we defined SS3 cells (similar to ID cells) as having densities between 1.091 and 1.105 g/mL, whereas SS4 cells
(similar to the most dense ID cells and all of the HD cells) have
densities greater than 1.105 g/mL. SS3 cells and SS4 cells differ in a
number of properties that support the contention that they should be
studied separately and may have different origins: SS3 cells have a
smaller percentage of ISCs (22.5% ± 12.8% v 70.2% ± 7.4%),5 a higher percentage of reticulocytes (6.3% ± 7.2% v 3.2% ± 5.8%),5 and a higher
percentage of HbF (8% ± 2% v 4% ± 2%).9
Moreover, Kaul et al5-7 have demonstrated that, whereas
both SS3 and SS4 cells are rheologically incompetent, SS3 cells show
the greatest change in rheological properties when oxygenated and
deoxygenated conditions are compared. That is, SS4 cells are
constitutively rigid, whereas SS3 cells rapidly become rigid due to HbS
polymer formation.
The dehydration of SS cells is thought to involve a
deoxygenation-induced increase in membrane permeability to monovalent alkali metals (Na+, K+) and divalent cations
(Ca2+, Mg2+) that is unique to SS cells
(reviewed in Canessa,8 Joiner,21 and Brugnara
and Tosteson22). The existence of a
Ca2+-dependent K+ channel
[K(Ca2+)]23-26 and a K:Cl cotransport (K:Cl)
pathway in red blood cells27-32 and particularly in SS red
blood cells33,34 is well established, but their relative
contributions to the formation of dense cells has remained
controversial.8,21,22 It has been suggested that
deoxygenation-induced stimulation of the K(Ca2+) results
secondarily and persistently in the activation of K:Cl cotransport by
virtue of the cellular acidification that accompanies cell
dehydration.26,35 Nevertheless, the relative importance and
individual effects of these pathways on dense SS cell formation are not
clearly understood.
Other lines of evidence suggest that the formation of dense dehydrated
SS cells may proceed by multiple pathways, some of which may be
dependent on cell age and HbF content. For example, the dense cell
population derived from whole SS blood contains both reticulocytes and
more mature, although still young, cells,26 indicating that
some SS cells become dense while very young, whereas other cells take
considerably longer to become dense. The presence of a rapid
dehydration pathway was first proposed by Bertles and Milner36 and later described in more detail by Bookchin et
al.26 Moreover, cells in the high density fraction (SS4)
contain a lower percentage of HbF than cells in the intermediate
density fraction (SS3).9 This difference is compatible with
a model in which SS4 cells are generated directly by a
polymerization-related mechanism and SS3 cells by a
polymerization-independent mechanism.
Fabry et al9 also demonstrated that only a fraction of SS2
cells have K:Cl volume regulation. More recently, Franco et al17 reported that, if SS cells are separated into very
young (transferrin-receptor positive [TfR+]) SS
reticulocytes and older cells (TfR), the
TfR+ cells have a lower HbF than TfR
cells, which they and others have attributed to a longer circulating lifespan of HbF-containing cells and the resultant enrichment in older,
high HbF cells. They also found TfR+ cells in the density
fraction defined as greater than 1.092 g/mL. That very young
reticulocytes can become dense implies a rapid path for dehydration and
their low level of HbF suggests that either their dehydration is
polymer-driven, because of the inhibitory effect of HbF on
polymerization, or that the older cells are HbF-rich, because these
cells are protected from polymer-driven destruction.17 Recently, Franco et al37 reported that, when unfractionated SS cells were subjected to cyclic oxygenation-deoxygenation (O-D) and
then subjected to density fractionation, the TfR+ cells had
undergone a larger density increase than the TfR
cells. They also demonstrated that, under conditions of cyclic O-D,
both K:Cl and K(Ca2+) pathways play a role in SS red blood
cell dehydration, which is in agreement with our previous
report.38 However, they did not analyze their data in terms
of different density classes of high-density cells.
Recent in vitro studies by others using repetitive O-D cycling to
generate dehydrated SS cells used conditions that favored ID cell
formation: short O-D cycling times (<6 hours) and buffer pH
7.4.25,37 Formation of HD cells in these studies was either negligible or not separately studied; thus, mechanisms by which HD
cells are formed in vivo could not be evaluated. This is important, because dense SS cells are heterogeneous in terms of their functional abnormalities and contribution to disease pathophysiology. Cells in the
ID cell fraction are more likely to be vaso-adhesive than HD cells,
which are more likely to be mechanically rigid and participate in
vaso-occlusive events.5,6
To test the hypothesis that different mechanisms may be involved in the
sickling-induced formation of HD and ID SS cells, we used a technique,
previously used by others,39-41 that simulates in vivo
sickling by exposing SS cells to repetitive in vitro O-D cycles. Unlike
some earlier studies, experimental conditions were selected to allow
for in vitro generation of both ID and HD cells from a density-defined
discoidal subpopulation (ND) of SS cells.
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MATERIALS AND METHODS |
Blood collection.
Blood was drawn from SS patients followed in the Day Hospital (Dr L. Benjamin, Director) or Hereditary Clinic (Dr H. Lachman, Director) of
the Bronx Comprehensive Sickle Cell Center (Bronx, NY) after informed
consent had been obtained. Diagnosis of homozygous  S
genotype was based on two types of electrophoresis (cellulose acetate,
borate buffer, pH 8.6, and agar, citrate buffer, pH 6.4), an
Mst II restriction digest gene analysis, and a solubility test. Patients with concomitant  -thalassemia and those transfused within 3 months were excluded.
Primary density fractionation.
To avoid the complication of following new dense cell formation in the
presence of pre-existing dense cells, all experiments were performed on
an ND cell fraction, prepared as described. SS blood was washed 3 times
in 10 mmol/L sodium phosphate, 140 mmol/L NaCl, pH 7.4 (PBS), and 0.5 mL of the washed cells were applied to a total volume of 6 mL of a
continuous isotonic density gradient, composed of Percoll-Larex UF
(Larex arabinogalactan; Larex International, St Paul, MN), and the
cells were separated by centrifugation as previously
described.19 The ND discocyte fraction (1.080 to 1.090 g/mL, similar to SS2) was isolated, as previously
described,19 and used as starting material for all of the
following experiments. Color instant Polaroid photographs and color transparencies (Polaroid Corp, Cambridge, MA) of
the density gradients were taken. To quantitate cells in the different density fractions, analytical gradients were run simultaneously using
0.05 mL packed red blood cells/6 mL gradient, and the color transparencies were scanned by laser densitometry.
In vitro O-D cycling.
One milliliter of approximately 40% hematocrit ND SS cells was added
to 14 mL of O-D cycling buffer in 50-mL Erlenmeyer flasks. O-D cycling
buffer typically contained (as final concentrations) 20 mmol/L sodium
HEPES, pH 7.4, 120 mmol/L sodium chloride, 0.9 mmol/L sodium phosphate,
4.5 mmol/L potassium chloride, 0.5 to 2 mmol/L calcium chloride (as
indicated), 0.9 mmol/L magnesium chloride, 10 mmol/L glucose, 0.2%
bovine serum albumin (BSA), 0.1 mg/mL streptomycin, and 100 U/mL
penicillin at 290 mOsm/kg. For experiments in which
Cl was excluded, all salts in the O-D cycling buffer
had nitrate (NO3) substituted for
Cl.
The flasks were sealed with rubber stoppers and continuously flushed
with alternating cycles of 5% CO2/balance air and 5% CO2/balance N2. Where buffer pH was 7.4, it was
adjusted after equilibration with 5% CO2. Where buffer pH
was 6.8, no adjustments were made and, in the absence of bicarbonate to
buffer the CO2, the resulting pH after equilibration with
CO2 was 6.8. Gas flow was set to approximately 30 mL/min.
O-D cycling was accomplished using a three-way valve and intervolameter
timer. A manifold was used to allow control and experimental conditions
to be tested simultaneously on the same sample. During the course of
our experiments, various times for the duration of the O and D
component of the O-D cycle were tested; they ranged from 5 minutes
(pO2 84 ± 4 mm Hg) to 15 minutes (pO2 118 ± 11 mm Hg) for the O cycle; for the D cycle, 10 and 15 minutes
were tested and found to yield the same value (pO2 22 ± 13 mm Hg). For some experiments, buffer pH was continuously monitored
using a needle pH electrode inserted into the sealed flask. The flasks
were shaken at 80 to 90 rpm at 37°C. After O-D cycling (or exposure
to continuous air or N2 equilibrated with 5%
CO2), 100% humidified oxygen was introduced into the
sealed flasks for an additional 30 minutes. Buffer pH at the end of the
oxygenation period was 7.2 to 7.3 and was similar for
Cl- and NO3-containing buffers. After 22 hours, buffer osmolality increased from 294 ± 3 to 310 ± 3 mOsm/kg. Hemolysis after 22 hours, measured as Hb released into the
buffer, was 2.3% ± 0.6% when the incubations took place at pH 7.4 and 5.6% ± 0.9% when the incubations took place at pH 6.8 (see
below for method). Hemolysis was not affected by increasing buffer
Ca2+ from 0.5 to 2 mmol/L or by using continuous gas
exposure versus O-D cycling.
Secondary density fractionation.
After oxygenation, the flasks were opened, and the cells collected by
centrifugation and washed three times in isotonic saline. The washed
cells were then separated a second time by analytical and preparative
density fractionation, using continuous Percoll-Larex gradients, as
described above.19 Photographs and transparencies of the
analytical gradients were taken. Density fractions were collected from
the preparative gradients by cutting at defined density locations to
yield low density (LD; <1.080 g/mL; MCHC, <32 g/dL), ND (1.080 to
1.090 g/mL; MCHC, 32 to 36 g/dL), ID (1.090 to 1.114 g/mL; MCHC, 36 to
46 g/dL), and HD (>1.114 g/mL; MCHC, >46 g/dL) cell fractions. This
separation is very similar, but not identical to, our previously
defined density populations SS1, SS2, SS3, and SS4.19 By
analogy to this earlier separation scheme, the majority of
reticulocytes are found in the LD and ND cell fractions.19
In some experiments, a single density cut was made at 1.10 g/mL (MCHC,
40 g/dL), resulting in two cell populations: a top fraction containing
lighter cells with densities less than 1.10 g/mL (MCHC, <40 g/dL) and
a bottom fraction containing the most severely dehydrated cells with
densities greater than 1.10 g/mL (MCHC, >40 g/dL). The reason for
this density cut was to increase the number of dense cells available
for this and other studies. Isolated density fractions were washed
three times in isotonic saline to remove the gradient mixture. Aliquots
of washed cells were fixed in glutaraldehyde for light and electron
microscopy, as described below.
ATP, Hb, and F-cell determinations.
For ATP, 0.8 mL of the O-D cycling mixture (at 0 hours and after 22 hours of O-D cycling) was added to 0.8 mL ice-cold 12% trichloroacetic
acid and vortexed to lyse the cells. The lysates were incubated on ice
for 5 minutes, quick frozen using a dry ice-ethanol bath, and stored
for less than 5 days at  80°C. Immediately before analysis
(all of the samples were analyzed on the same day), the lysates were
thawed and the supernatants were collected by microfuge. ATP was
assayed using Sigma kit 366-UV (Sigma, St Louis, MO),
according to the manufacturer and expressed as micromoles of ATP per
gram of Hb. Hb was determined in 0.05 mL of the O-D cycling mixture
using Drabkins reagent and compared with an Hb standard, according to
Sigma kit 525 (Sigma).
For hemolysis determinations, Hb was determined in 0.5 mL of the O-D
cycling buffer (after centrifuging out the intact red blood cells)
using Drabkins reagent, as described above, and compared with a 100%
hemolysis standard.
F cells were measured by fluorescence-activated cell sorting
(FACS) analysis using PBS-washed red blood cells derived
from whole blood or Percoll-Larex density fractions before and after O-D cycling using a protocol performed and developed by Wallac/Isolab, Inc (Akron, OH). Briefly, after washing red blood cells from blood or
Percoll-Larex density fractions at least four times with PBS, the cell
pellet was resuspended in an approximately equal volume of PBS. For
fixation, 10 µL of the cell suspension was added to 1 mL of PBS/4%
formaldehyde (Fisher Scientific, Pittsburgh, PA) for 1 hour at room
temperature. PBS (0.25 mL)/0.05% glutaraldehyde (Fisher Scientific,
Pittsburgh, PA) was added, and the cells were mixed for an additional
30 seconds and then immediately pelleted by centrifugation for 5 minutes at 150g. A brief exposure to glutaraldehyde was found
to aid in preserving cell morphology; however, care must be taken to
limit exposure to the glutaraldehyde to 30 seconds plus the 5 minutes
of centrifugation time. The fixed cells were then washed in PBS,
centrifuged, resuspended in 0.25 mL of PBS/5% nonfat dry milk, mixed
gently for 10 minutes at room temperature, and centrifuged. For
permeabilization, the cell pellet was resuspended in 0.5 mL of 0.01%
Triton X-100 (Sigma/Aldrich, Milwaukee, WI) in PBS/0.1% BSA and
aliquoted into two 0.1-mL fractions. One aliquot received fluorescein
isothiocyanate (FITC)-conjugated monoclonal antibody to HbF
(Wallac/Isolab, Inc; affinity purified, 1 µg/0.1 mL cell suspension),
whereas the other aliquot received an isotypic control. The cells were
mixed gently for 30 minutes at room temperature, centrifuged, washed in
PBS, pelleted, and resuspended in PBS. A minimum of 10,000 cells were
analyzed using an Epics XL flow cytometer (Coulter, Hialeah, FL). For
some experiments, red blood cell HbF was determined in hemolysates by
electrophoresis on citrate agar.9
Ion transport inhibitors.
In some experiments, pharmacologic inhibitors of the K:Cl cotransporter
or K(Ca2+) were added. To inhibit K:Cl cotransport, final
concentrations of 0.5 µmol/L okadaic acid (ammonium salt; Research
Biochemical International, Natick, MA; stock solution at 0.1 mmol/L in
dimethyl sulfoxide [DMSO]) and 0.1 mmol/L
[(dihydroindenyl)oxy]alkanoic acid (DIOA; Research Biochemical
International, Natick, MA; stock solution at 50 mmol/L in DMSO) were
used. To inhibit K(Ca2+), final concentrations of 10 µmol/L clotrimazole (Sigma; 10 mmol/L stock solution in DMSO) and 0.1 µmol/L charybdotoxin (Bachem Bioscience, Philadelphia, PA; stock
solution at 0.15 mmol/L in water) were used. These experiments also
contained 0.1 mmol/L ouabain to inhibit the
Na+-K+-ATPase (Sigma; 10 mmol/L stock in water)
and 10 µmol/L bumetanide to inhibit the
Na+-K+-2Cl cotransporter (Sigma;
10 mmol/L stock in DMSO).
Microscopy.
Cells (0.05 mL packed cells) were fixed for light and electron
microscopy in 2 mL ice-cold 2% glutaraldehyde in PBS, pH 7.4. After
incubation for 1 hour on ice, the cells were collected by centrifugation and washed three times in PBS. Fixed cells were used to
make smears on slides that were stained with Giemsa stain in an
automated slide stainer. Cells were examined under 1,000× light
microscopy and photographs were taken of several fields. ISC counts
were made from the photographs. Morphologically, ISCs were strictly
defined as cells whose length was  2× width and having only two
points at opposite ends of the cell after oxygenation for 30 minutes in
100% oxygen. The same investigator performed all of the ISC counts.
Statistical analysis.
Values are presented as the mean ± 1 SD. Statistical analysis was
performed using SigmaStat for Windows (ver 1.0; Jandel Scientific, San
Rafael, CA). The probability of the null hypothesis was calculated using the paired or unpaired t-test (two-tailed), as
appropriate.
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RESULTS |
Effect of buffer pH and number of O-D cycles on dense cells formed from
ND SS cells by O-D cycling.
O-D cycled cells were fixed in glutaraldehyde in both the oxygenated
(O) and deoxygenated (D) stages of the O-D cycle. Cells fixed in the D
stage were greater than 90% sickle shaped, with many cells exhibiting
multiple projections, whereas cells fixed in the O stage were
predominately biconcave discs, demonstrating that our O-D cycling
protocol resulted in near complete cell sickling-unsickling during each
O-D cycle (data not shown). This was true for O stages that ranged from
3 to 15 minutes and for D stages that ranged from 8 to 15 minutes.
O-D cycling SS ND cells for 22 hours (88 cycles of 5 minutes O and 10 minutes D) in buffer containing physiologic levels of sodium,
K+, and Cl and 0.5 mmol/L
Ca2+ in 5% CO2-equilibrated buffer at pH 6.8 resulted in both ID cell (density, 1.090 to 1.114 g/mL) and HD cell
(density, >1.114 g/mL) formation (Fig
1A). In contrast, O-D cycling at pH 7.4 in 5%
CO2-equilibrated buffer did not result in generation of HD
cells and only small numbers of cells that were more dense than the
parent ND cells were formed (Fig 1A).
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| Fig 1.
Formation of dense SS cells by in vitro O-D cycling. SS
ND (1.080 to 1.090 g/mL) cells were isolated from SS whole blood by
density gradient centrifugation using continuous density isotonic
Percoll-Larex (Stractan) gradients with physiologic concentrations of
Na+, K+, Mg2+, and
Cl. The ND cells were subject to O-D cycling in sealed
flasks containing a bicarbonate-free HEPES buffer at pH 7.4 using 5%
CO2/balance air (duration, 5 minutes) and 5%
CO2/balance N2 (duration, 10 minutes) for the
oxy (O) and deoxy (D) stages, respectively. Buffer pH was adjusted to
7.4 after equilibration with 5% CO2. Without adjustment,
in the absence of bicarbonate and after equilibration with 5%
CO2, the resulting buffer pH was 6.8. Experiments using
continuous deoxygenation or continuous oxygenation also used 5%
CO2-containing gasses. After O-D cycling (or continuous air
or N2 exposure) the flasks were flushed under sealed
conditions with 100% oxygen for 30 minutes, followed by collection of
the cells and a second density separation, as described above. Color
transparencies of analytical gradients were taken and scanned by laser
densitometry to quantitate the percentage of cells in each density
fraction. Cell density fractions were defined as follows: LD (<1.080
g/mL), ND (1.080 to 1.090 g/mL), ID (1.090 to 1.114 g/mL), and HD
(>1.114 g/mL). Results shown are representative of from 2 to 15 experiments and are paired results using a single SS patient in all
arms of the experiment. (A) Effect of pH on dense cells formed after 22 hours of O-D cycling. (B) Time course of dense cells formed by
continuous deoxygenation (5% CO2/balance N2)
or O-D cycling at pH 6.8. (C) Dense cells formed after 22 hours of
continuous oxygenation (5% CO2/balance air) or O-D cycling
at pH 6.8. (D) Effect of ATP depletion on dense cell formation by O-D
cycling. The O-D cycling buffer contained either 10 mmol/L glucose
(Gluc) or 10 mmol/L 2-deoxyglucose (D-Gluc). The final concentration of
Ca2+ in the buffer was 1.5 mmol/L (for A and D) or 0.5 mmol/L (for B and C). B, density marker beads (Amersham Pharmacia
Biotech, Piscataway, NJ).
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At pH 6.8, ID cells first became apparent after 3 hours of continuous
deoxygenation or O-D cycling (Fig 1B). At 8 hours, there was
considerable ID cell formation that was increased in continuously deoxygenated cells compared with O-D cycled cells. HD cell formation was not significant until 8 hours for both continuously deoxygenated and O-D cycled cells. After 22 hours, greater than 50% of the parent
SS ND cells were found in the dense cell fractions (ID + HD; Fig 1A and
B are representative, and data from 15 other SS patients are summarized
in Fig 5, 22-hour chloride data).
SS patients were heterogeneous in the number of ID and HD cells formed
by continuous deoxygenation or O-D cycling (data not shown). We noted
that ID cell formation increased substantially when
CO2-free gasses were used (data not shown). This may be due to the recently described inhibition of K:Cl activity by
HCO3 at plasma levels (~24
mmol/L)42; our buffer level of
HCO3 after CO2-equilibration
is approximately 24 mmol/L at pH 7.4, but only approximately 6 mmol/L
at pH 6.8 (see Discussion).
Five percent to 15% of the cells that had been O-D cycled at pH 6.8 for 22 hours in 5% CO2-equilibrated buffer (containing 0.5 mmol/L Ca2+) and then oxygenated with 100% oxygen for 30 minutes had a morphology closely resembling endogenous ISCs, ie,
elongated cells with single projections at each end. Increasing
oxygenation time to 1 hour or exposing the oxygenated cells to carbon
monoxide (1 hour at 4°C) did not reduce ISC formation or change ISC
morphology (data not shown). Moreover, electron micrographs of thin
sections confirmed the absence of HbS polymer, establishing their
classification as ISCs (data not shown). ISCs were more numerous in the
most dehydrated bottom density cell fraction (density, >1.10 g/mL; MCHC, >40 g/dL) compared with lighter cells (density, <1.10 g/mL; Fig 2D and C, respectively).
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| Fig 2.
Morphology of newly formed dense cells. After 22 hours at
pH 6.8 in 5% CO2-equilibrated buffer, as in Fig 1 (for
B-F), cells were oxygenated for 30 minutes with 100% oxygen, fixed in
glutaraldehyde, and examined by light microscopy. (A) Uncycled parent
SS ND cells; (B) SS ND cells exposed to continuous oxygenation for 22 hours; (C) top density cell fraction (<1.10 g/mL) formed after 22 hours of O-D cycling; (D) bottom density cell fraction (>1.10 g/mL)
formed after 22 hours of O-D cycling; (E) top density cell fraction
(<1.10 g/mL) formed after 22 hours of continuous deoxygenation; and
(F) bottom density cell fraction (>1.10 g/mL) formed after 22 hours
of continuous deoxygenation.
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Exposure of ND SS cells to continuous oxygenation with air for 22 hours
at pH 6.8 with 5% CO2 equilibrated buffer resulted in less
than 10% dense cell formation (ID + HD), much less than that formed by
O-D cycling (Fig 1C), and most of the continuously oxygenated cells had
a morphology similar to uncycled parent ND cells (Fig 2B and A,
respectively). Many of the continuously deoxygenated cells had multiple
projections, clearly different from the morphology of endogenous ISCs.
These abnormally shaped cells were more common in the dehydrated cell
fraction (density, >1.10 g/mL) compared with the lighter cells
(density, <1.10 g/mL; Fig 2F and E, respectively). Hematologically
normal AA cells exposed to either 22 hours of O-D cycling or continuous
deoxygenation at pH 6.8 resulted in less than 1% dense cell formation,
and cell morphology was normal (data not shown).
Effect of ATP depletion on dense SS cells formed by O-D cycling.
To evaluate the role of ATP depletion on O-D cycling-induced dense cell
formation, SS ND cells were O-D cycled for 22 hours at pH 6.8 in 5%
CO2-equilibrated buffers, containing either 10 mmol/L
glucose or 10 mmol/L 2-deoxyglucose, a nonmetabolizable carbohydrate
used by others to ATP-deplete red blood cells.43 As shown
in Fig 1D, O-D cycling for 22 hours in the presence of 2-deoxyglucose
resulted in a large increase in the formation of HD cells and a
decrease in the formation of ID cells compared with O-D cycling in the
presence of glucose. In six SS patients in whom ATP levels before and
after 22 hours of O-D cycling at pH 6.8 in the typical O-D cycling
buffer (containing 10 mmol/L glucose) were measured, ATP levels after
O-D cycling were heterogeneous, with no appreciable ATP loss in three
SS patients and decreases of between 26% and 58% in three SS patients
(Fig 3). However, in all six SS patients,
and in every SS patient we have studied, there was formation of both ID
and HD cells under these conditions.
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| Fig 3.
ATP levels after 22 hours of O-D cycling SS ND cells at
pH 6.8. Six paired SS patient samples were tested before and after O-D
cycling as described in Fig 1. The cells collected after O-D cycling
represented all cells, that is, before density separation. ATP levels
are expressed as micromoles per gram of Hb.
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|
F-cell content of ID and HD cells formed by O-D cycling.
The F-cell content of unfractionated red blood cells, ND cells derived
from whole blood, and ID and HD cells formed from the ND cells after 22 hours of O-D cycling at pH 6.8 in 5% CO2-containing gasses
was determined by FACS analysis in four SS patients
(Fig 4). ID cells formed after 22 hours of
O-D cycling at pH 6.8 contained a percentage of F cells that was only
slightly lower than the ND cells from which they were derived (ID/ND = 0.76 to 0.95). In contrast, HD cells formed after 22 hours of O-D
cycling at pH 6.8 contained a much lower percentage of F cells compared
with the ND cells from which they were derived (HD/ND = 0.42 to 0.74).
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| Fig 4.
The percentage of F cells in unfractionated, ND uncycled,
and ID and HD cells formed after 22 hours of O-D cycling at pH 6.8. SS
ND cells derived from SS whole blood (WB) were O-D cycled for 22 hours
at pH 6.8, and the newly formed dense cells were separated into ID and
HD cells using continuous density gradients, as in Fig 1. Results are
shown for four SS patients.
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Role of the K:Cl cotransport in dense SS cell formation.
To evaluate the role of K:Cl cotransport in O-D cycling-induced dense
SS cell formation, Cl in the O-D cycling buffer was
replaced by NO3 to inhibit this pathway.
Experiments were conducted using 15 paired samples of ND SS cells O-D
cycled for 22 hours at pH 6.8 in 5% CO2-containing gasses.
As shown in Fig 5, O-D cycling in buffer
containing Cl resulted in the formation of 35% ± 17% ID cells and 18% ± 11% HD cells. Substitution of
NO3 for Cl reduced ID
cell formation to 15% ± 9% (P < .0004)
without significantly affecting HD cell formation, along with a
concomitant increase in parent ND cells compared with O-D cycling in
Cl (P < .0071).
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| Fig 5.
Effect of chloride or nitrate on dense cells formed by
O-D cycling. SS ND cells were O-D cycled for 22 hours at pH 6.8 in
buffer containing chloride or nitrate, each with 0.5 mmol/L
Ca2+. After O-D cycling, the cells were reapplied to
density gradients and LD, ND, ID, and HD fractions were quantitated as
described and defined in Fig 1. Results are presented for 15 SS
patients where paired data were available. p calculated using
the paired t-test (two-tailed).
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Effect of calcium on dense SS cell formation.
HD cell formation at pH 6.8 in 5% CO2-containing gasses
was enhanced by the presence of Ca2+ in the cycling buffer,
but the requirement for Ca2+ was not absolute, because
substitution of Ca2+ with EGTA reduced, but did not
completely prevent, HD cell formation (Table 1). Consistent with this finding,
increasing the concentration of Ca2+ in the O-D cycling
buffer from 0.5 to 2 mmol/L increased approximately twofold HD cell
formation in two SS patients (Table 2).
Effect of ion transport inhibitors on dense SS cell formation.
Ouabain (100 µmol/L) and bumetanide (10 µmol/L), inhibitors of the
Na+-K+-ATPase and
Na+-K+-2Cl cotransporter,
respectively, had no detectable effect on 22-hour O-D cycling-induced
dense cell formation at pH 6.8 in 5% CO2-containing gasses
in the absence of other inhibitors (data not shown). Inhibitors of K:Cl
cotransport, DIOA (100 µmol/L) and okadaic acid (0.5 µmol/L), inhibited by 84% ± 8% (n = 4) the formation of ID cells but did not appreciably inhibit the formation of HD cells. This was observed in
SS patients who produced large numbers of ID cells (G.E. in Fig 6A is representative), intermediate
numbers of ID cells (A.C. in Fig 6A is representative), and low numbers
of ID cells (M.R. in Fig 6A is representative). In some SS patients we
noted a small increase in the number of HD cells formed in the presence
of DIOA, but not okadaic acid. In contrast, inhibitors of
K(Ca2+), charybdotoxin (0.1 µmol/L), and clotrimazole (10 µmol/L) inhibited by 42% ± 28% (n = 5) the formation of HD
cells without appreciably inhibiting ID cell formation. This was
observed in SS patients who produced large numbers of HD cells (J.C. in
Fig 6B is representative), intermediate numbers of HD cells (M.M. in
Fig 6B is representative), and low numbers of HD cells (M.R. in Fig 6B
is representative). Figure 6C shows the results of an experiment in
which both K:Cl and K(Ca2+) inhibitors were tested alone or
together on the same SS patient. Consistent with results shown in Fig
6A and B, DIOA primarily inhibits ID cell formation and clotrimazole
primarily inhibits HD cell formation. Combined use of both DIOA and
clotrimazole nearly abolished ID and HD cell formation. The small
number of dense cells that were formed in the presence of these
inhibitors were irregularly shaped, suggesting that cell damage had
occurred (data not shown).
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| Fig 6.
Effect of ion transport inhibitors on dense cell
formation by O-D cycling. SS ND cells were O-D cycled for 22 hours at
pH 6.8 and density separated into LD, ND, ID, and HD fractions, and the
percentage of cells in each fraction was quantitated as described and
defined in Fig 1. All experiments contained ouabain (100 µmol/L final
concentration) and bumetanide (10 µmol/L final concentration) to
inhibit the Na+-K+-ATPase and
Na+-K+-2Cl cotransporter,
respectively. (A) Inhibitors of K:Cl cotransport, DIOA (100 µmol/L
final concentration), and okadaic acid (OKA; 0.5 µmol/L final
concentration) primarily inhibited ID cell formation (1.090 to 1.114 g/mL). (B) Inhibitors of K(Ca2+), charybdotoxin (CTX; 0.1 µmol/L final concentration), and clotrimazole (CLT; 10 µmol/L final
concentration) inhibited primarily HD cell formation (>1.114 g/mL).
(C) Combined use of K:Cl cotransport inhibitor (DIOA) and
K(Ca2+) inhibitor (CLT) in the same SS patient on ID and
HD cell formation. Inhibitor concentrations are as described above. A
manifold was used to allow for the simultaneous O-D cycling of all four
test conditions.
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DISCUSSION |
SS red blood cells in sickle cell blood are heterogeneously dehydrated,
with some cells becoming HD (defined as >1.114 g/mL; MCHC, >46
g/dL; similar to SS4) and others dehydrating to ID (defined as 1.090 to
1.114 g/mL; MCHC, 36 to 46 g/dL; similar to SS3; reviewed in Nagel and
Fabry44). Although it is clear that both the
K(Ca2+) and the K:Cl cotransport pathways contribute to SS
red blood cell dehydration, it is not known to what extent each pathway is involved in the generation of different classes of dense SS cells.
To test the hypothesis that ID and HD cells are generated by separate
pathways, we adapted an in vitro O-D cycling protocol to generate ID
and HD cells from ND SS discocytes. The O-D cycling protocol was
adapted from earlier protocols originally described by Ohnishi et
al39 and modified by others40,41 and has been used to define mechanisms involved in sickling-induced ISC and dense
cell formation and SS red blood cell rheologic abnormalities. Our
adaptation was to perform the O-D cycling at pH 6.8 (in buffer equilibrated with 5% CO2) at which activity of K:Cl
cotransport is maximal33 and sickling is favored. In
addition, we started with a relatively homogeneous ND cell population
that precludes the problems of trying to measure newly formed dense
cells in a mixed population containing preformed dense cells that may
also change their distribution during O-D cycling. Because the ND cell fraction (similar to SS2) is enriched in young cells and reticulocytes (reviewed in Nagel and Fabry44), it is likely that this
fraction is physiologically relevant for the study of dense SS cell
formation in vivo. However, we recognize that our ND fraction is a
selected cell fraction and that it does not contain cells that became
dense in vivo. Thus, it is possible that we may not be studying cells that have the greatest tendency to become dense. Unfortunately, it is
impossible to test this experimentally, because one cannot study dense
cell formation in vitro using preformed dense cells.
With our O-D cycling protocol (physiologic levels of Na+,
K+, Mg2+, and Cl; 0.5 mmol/L
to 2 mmol/L Ca2+; and equilibration with
5%CO2, at pH 6.8), greater than 50% of the parent ND SS
cells dehydrated (they are recovered in the ID and HD cell fractions)
after O-D cycling for 22 hours, and some cells were morphologically
similar to endogenous ISCs. These conditions were selected because,
when shorter O-D cycling times (<8 hours) or buffer pH 7.4, in the
absence of CO2, were used by others,25,37 the
yield of newly formed high-density cells (>1.103 g/mL, as defined in
Brugnara et al25 and Franco et al37) was low.
O-D cycling ND SS cells for 22 hours at pH 7.4 (in 5%
CO2-equilibrated buffer) resulted in a small amount of ID
cell formation that was seen in some but not all SS patients, and there
was no appreciable HD cell formation in any SS patient. In contrast, O-D cycling at pH 6.8 (in 5% CO2-equilibrated buffer) for
22 hours resulted in approximately 20% of the parent ND cells
dehydrating to the HD cell fraction. Exposure of ND SS cells to
continuous air for 22 hours at pH 6.8 (in 5%
CO2-equilibrated buffer) resulted in less than 10% dense
cell (ID + HD) formation, substantially less than that formed by O-D
cycling (>50%). This supports the contention that the formation of
HbS polymer is essential for the generation of at least some types of
dense cells in vitro. HD cells were also formed by exposure of ND cells
to continuous deoxygenation (N2/5% CO2) for 22 hours at pH 6.8; indeed, continuous deoxygenation at pH 6.8 promoted HD
cell formation to an even greater extent than did O-D cycling. This
suggests that continuous deoxygenation at pH 6.8 stimulates
K+ loss by the K(Ca2+) pathway. Our results are
consistent with those of Franco et al,37 who reported that
continuous deoxygenation of TfR+ SS cells resulted in
density increases that were inhibited by removal of Ca2+
but not by replacement of Cl with
NO3, suggesting that dense cells formed
by continuous deoxygenation (at pH 7.4) proceeded by a
Ca2+-dependent, Cl-independent pathway,
presumably the K(Ca2+) pathway, and did not require the
K:Cl cotransport pathway. Alternatively, because ATP depletion
accelerates HD cell formation, it is possible that ATP levels are
affected differently depending on whether the cells are repetitively
sickled or continuously deoxygenated. Another possibility is that
continuous deoxygenation inhibits K:Cl cotransport activity. This
possibility is consistent with the results of Franco et
al,37 who reported that cyclic O-D but not continuous
deoxygenation results in an increase in SS red blood cell density
mediated by a Cl-dependent pathway, presumably the
K:Cl cotransporter.
Consistent with the hypothesis that the K(Ca2+) pathway is
principally involved in HD cell formation (by O-D cycling at pH 6.8), addition of the Ca2+ chelator EGTA to the O-D cycling
buffer inhibited, but did not prevent, HD cell formation, and
increasing Ca2+ in the O-D cycling buffer from 0.5 to 2 mmol/L increased HD cell formation without affecting ID cell formation.
ID cells formed after 22 hours of O-D cycling at pH 6.8 contained a
percentage of F cells that was only slightly smaller than the ND cells
from which they were derived (ID/ND = 0.76 to 0.95). In contrast, HD
cells formed after 22 hours of O-D cycling at pH 6.8 contained a much
smaller percentage of F cells compared with the ND cells from which
they were derived (HD/ND = 0.42 to 0.74). This result is consistent
with earlier findings that the most dense circulating SS cells had the
lowest HbF content9,36 and further suggests that the
formation of the most severely dehydrated cells (HD cells) is more
dependent on HbS polymerization than is the formation of ID cells.
These results are also consistent with our contention that ID and HD
cells can be formed by independent mechanisms.
Franco et al17 reported that the HbF content of very young
transferrin-receptor positive (TfR+) circulating SS cells
was negligible ( 0.6%) in dense cells (>1.092 g/mL, equivalent to
the densest 50% of our ID fraction and all of our HD cell fraction)
compared with TfR+ light cells (<1.092 g/mL), implying
that HbF protects against cell dehydration in this very young
TfR+ population. Thus, it is likely that HbF is protective
against cell dehydration both in the youngest and in the more mature, albeit still young SS cells that are destined to become the most severely dehydrated cells. Our results are also consistent with recent
data of Franco et al,45 who found that HbF is an important determinant in the formation of high-density (density, >1.094 g/mL)
TfR+ SS reticulocytes, but does not mediate the formation
of TfR+ moderate density (density, 1.083 to 1.094 g/mL) SS
reticulocytes. This is also in agreement with our previous conclusion
that both F reticulocytes and non-F reticulocytes have a similar K:Cl
driven response to hypotonic stimulation.9 Together, these
results suggest that sickling is not required to activate the K:Cl
cotransporter or form moderate density cells, but does not rule out the
possibility that sickling may enhance K:Cl cotransport activity and
increase ID cell formation. Regardless, the conclusions of Franco et
al45 and our conclusions from the results presented here
are in agreement that different mechanisms are likely to be involved in
the formation of different classes of dense young SS cells.
There are several arguments to support O-D cycling at pH 6.8 for 22 hours as pathophysiologically relevant:
(1) Although blood pH approaching 6.8 is not encountered in the general
circulation, acid pH is found in regions of the spleen,46 kidney,47 and in situations in which blood stasis occurs,
such as vessels which are partially vaso-occluded for short periods of
time.48 The final acidity depends in part on the length of stasis and the extent of deoxygenation. In this regard, SS patients are
in particular danger of long periods of stasis in organs with sinusoids
and in localized regions of the general circulation in which frequent
transient vaso-occlusions are likely.49
(2) This protocol generates both ID and HD cells (Fig 1) and better
represents than other methods the actual pattern of dense cells found
endogenously in SS whole blood.19
(3) Some of the O-D cycling-induced dense cells are morphologically
similar to endogenous ISCs (Fig 2), whereas dense cells resulting from
continuous deoxygenation contain multiple projections clearly
different from endogenous ISCs.
(4) HbF protects against the formation of the most severely dehydrated
HD cells generated by O-D cycling (Fig 4), consistent with the known
protective effect of HbF on in vivo dense cell formation (reviewed in
Nagel and Fabry44).
Hence, it is reasonable to contend that O-D cycling for 22 hours at pH
6.8 simulates dense cell formation in vivo, validating this approach.
ATP depletion is a potential complication in our studies using 22-hour
incubations, because, in the presence of Ca2+, ATP
depletion is associated with cell K+ loss and
dehydration.24 We find that, in the presence of 10 mmol/L
glucose, ATP loss was heterogeneous, with no significant loss for some
samples after 22 hours and decreases of between 26% and 58% in other
samples. That dense ID and HD cells were formed in all cases suggests
that ATP depletion is not required for ID and HD cell formation.
Substitution of 2-deoxyglucose, a nonmetabolizable
carbohydrate43, for glucose in the O-D cycling buffer
greatly stimulated HD cell formation and slightly inhibited ID cell
formation after 22 hours. These data are consistent with multiple
pathways being involved in the formation of dense SS cells and are
consistent with the known metabolic dependence of K:Cl cotransport
activity (reviewed in Lauf et al50). This conclusion is
also in accordance with the observation that ISCs are ATP depleted in
vivo51 and with the known stimulation of
K(Ca2+) by ATP-depletion in the presence of
Ca2+.52
It is unlikely that K:Cl cotransport is involved in generating the HD
cells in the ATP-depleted population, because increased free
Mg2+ resulting from decreased ATP:Hb ratios would be
expected to result in K:Cl cotransport inhibition,53 thus
supporting our argument that the most severely dehydrated HD cells are
more likely formed by the K(Ca2+) pathway (also see below).
Our experiments were performed using gasses containing 5%
CO2. It can be calculated that the
HCO3 concentration at equilibration with
dissolved CO2 at pH 6.8 is approximately 6 mmol/L and at pH
7.4 is approximately 24 mmol/L [log10(HCO3) = pH + log10(pCO2) 7.604 at 37°C, where
pCO2 at 5% CO2 = 38mm Hg]. It was recently
shown that SS red blood cell K:Cl cotransport activity is strongly
inhibited by physiologic levels of HCO3
found in plasma at pH 7.4 (~24 mmol/L),42 and this may
explain why we found little formation of ID cells at pH 7.4 using 5%
CO2-equilibrated buffer, whereas we found significantly
more ID cell formation at pH 7.4 using CO2-free buffer
(data not shown).
To study the contribution of the different transport pathways involved
in dense cell formation, we first defined the requirement for
Cl (indispensable for K:Cl cotransport in human red
blood cells) in O-D cycling-induced dense cell formation. Whereas
NO3 has been reported to bind Hb and
alter red blood cell volume,54 the effect is too small
(<10%) in human red blood cells to be of major concern, and the use
of other anions would prevent us from comparing results with a large
body of data previously collected using
NO3 to inhibit human red blood cell K:Cl
cotransport.
SS ND cells were O-D cycled in the presence of Cl or
NO3 (and 0.5 mmol/L Ca2+) at
pH 6.8. In Cl, both ID and HD cells were generated;
with NO3, there was a 43% reduction of
ID cells without significantly affecting HD cell formation. This
demonstrates that both a Cl-dependent and a
Cl-independent pathway are present for
sickling-induced SS cell dehydration from ND cells. We also noted
considerable individual to individual variability in the extent of ID + HD cell formation, both in the Cl and
NO3-O-D cycled experiments, suggesting
that the generation of ID + HD cells is subject to genetic or
epigenetic regulation and is consistent with previous observations of
variability in ion transport activity between SS
patients.8,9,53,55,56 It is likely that most of the K:Cl
cotransport activation occurs during the oxy stage of the O-D cycle,
because deoxygenation partially inhibits K:Cl cotransport
activity.53
The effect of ion transport inhibitors further addresses the issue of
the potential differences in the mechanisms of ID versus HD cell
formation. Both ID and HD cells are formed by O-D cycling in the
presence of Cl |
Top
Abstract
Introduction