Blood, 15 March 2001, Vol. 97, No. 6, pp. 1869-1875
RED CELLS
Membrane instability in late-stage erythropoiesis
Richard E. Waugh,
Athanassios Mantalaris,
Richard
G. Bauserman,
William C. Hwang, and
J. H. David Wu
From the Department of Pharmacology and Physiology,
University of Rochester School of Medicine and Dentistry and Department
of Chemical Engineering, University of Rochester, Rochester, New York.
 |
Abstract |
During maturation of the red blood cell (RBC) from the nucleated
normoblast stage to the mature biconcave discocyte, both the structure
and mechanical properties of the cell undergo radical changes. The
development of the mechanical stability of the membrane reflects
underlying changes in the organization of membrane-associated cytoskeletal proteins, and so provides an assessment of the time course
of the development of membrane structural organization. Membrane
stability in maturing erythrocytes was assessed by measuring forces
required to form thin, tubular, lipid strands (tethers) from the
surfaces of mononuclear cells obtained from fresh human marrow samples,
marrow reticulocytes, circulating reticulocytes, and mature
erythrocytes. Cells were biotinylated and manipulated with a
micropipette to form an adhesive contact with a glass microcantilever, which gave a measure of the tethering force. The cell was withdrawn at
controlled velocity and aspiration pressure to form a tether from the
cell surface. The mean force required to form tethers from marrow
reticulocytes and normoblasts was 27 ± 9 pN, compared to
54 ± 14 pN for mature cells. The energy of dissociation of the bilayer from the underlying skeleton increases 4-fold between the
marrow reticulocyte stage and the mature cell, demonstrating that the
mechanical stability of the membrane is not completely established
until the very last stages of RBC maturation.
(Blood. 2001;97:1869-1875)
© 2001 by The American Society of Hematology.
| |
Introduction |
During the last stages of maturation of
red blood cells (RBCs), dramatic changes occur in the
structure and organization within the cell. The cell loses its nucleus,
surface molecules are shed in small vesicles, and the final
surface-to-volume ratio of the cell is established.1-3
During this time, proteins that will eventually form the
membrane-associated cytoskeleton (membrane skeleton) are synthesized
and assembled at the intracellular surface of the plasma
membrane.4 The time course over which these protein assemblies become functionally viable is of interest, particularly with
regard to hemolytic anemia and the early release of cells during
hemorrhagic crisis, and could be important in designing methods for
production of erythrocytes in vitro.
The function of the assembled membrane skeleton is
fundamentally mechanical, and therefore, studies of membrane mechanical properties in maturing cells provide the most direct assessment of the
development of the functional viability of the skeleton during
maturation. Early studies of membrane properties of both murine and
human reticulocytes indicated increased membrane stiffness (shear
rigidity) in those membranes.5 This increased rigidity has
been confirmed subsequently both by micropipette6 and cell deformation in shear (ektacytometry).7 In the latter
study, evidence was also obtained that, despite increased mechanical stiffness, membranes of immature cells were less mechanically stable
than their mature counterparts, as indicated by fragmentation of cells
in fluid shear and in micropipette aspiration studies. The structural
events associated with membrane fragmentation may involve either
"tearing" of the membrane skeleton, or separation of the
membrane bilayer from the underlying skeleton. Early
fragmentation studies using ektacytometry showed a correlation between
decreased membrane stability and chemical abnormalities related to
lateral associations within the membrane skeleton, suggesting that the fragmentation involved mechanical failure of the skeleton
itself.8 More recent studies using fluorescence imaging of
micropipette-deformed cells showed that fragmentation in micropipettes
involves lateral segregation of membrane components and separation of
the lipid bilayer from the underlying skeleton.9
Thus, fragmentation may occur as a result of different
underlying mechanisms. An alternative to fragmentation methods for
assessing membrane instability is the formation of membrane
strands (tethers) from the surfaces of cells.10-12 This
approach provides a direct and quantitative measure of the energy
required to separate the membrane bilayer from the underlying
membrane-associated cytoskeleton.13,14 In the present
report we have applied this approach to demonstrate that the
instability observed in immature RBCs lies in the tightness of the
association between membrane bilayer and the underlying membrane
skeleton, and we provide a quantitative measure of the degree of
instability in terms of the work (energy) required to separate lipid
bilayer from the surfaces of cells at different stages of maturation.
| |
Materials and methods |
Cell source and separation procedure
Circulating cells.
Normal mature human erythrocytes were obtained by venipuncture from
healthy donors after informed consent according to
the University of Rochester's Research Subjects Review Board.
Heparin was used as anticoagulant. The reticulocytes were isolated by positive selection using magnetic beads coated with antibodies against
transferrin receptor (CD71-M450, Dynal, Lake Success, NY. See Brun and
coworkers.15). White blood cells were first removed by
filtration through a cellulose column (Sigma Cell type-50, Sigma
Chemical, St Louis, MO). A gentle negative pressure was applied to
facilitate the filtration. A volume of 250 µL filtered whole blood
was mixed with 25 µL magnetic beads on a rocker at room temperature
for 1 hour. The remaining filtered whole blood was spun at 2500 rpm for
10 minutes to isolate plasma (centrifuge model HN-SII, International
Equipment, Needham Heights, MA). The reticulocyte-bead mixture was
washed 2 times in HEPES buffered saline (130 mM NaCl, 10 mM HEPES, plus
2.0 g/L bovine serum albumin [BSA], pH 7.3 ± 0.1) using a
magnet to separate cells adhering to beads from the rest of the
population. After washing, the beads were separated from the cells by
incubation in plasma (which contains soluble transferrin receptor that
competes with the cells for binding sites on the beads) on a rocker at
room temperature for 1 hour.
Marrow cells.
Human bone marrow cells were aspirated from the iliac crest of healthy
donors after informed consent according to the University of
Rochester's Research Subjects Review Board. The total marrow cells
were diluted 1:1 with McCoy 5A medium (Gibco, Grand Island, NY) and
layered onto Ficoll-Paque (1.077 g/mL; Pharmacia, Piscataway, NJ).
Mononuclear cells (MNCs) from the interface band were collected after
centrifugation at 300g for 30 minutes at room temperature. The MNCs were washed twice in McCoy 5A medium and then resuspended at a
cell density of 1 × 106 cells/mL in filter-sterilized
phosphate-buffered saline (PBS; Gibco) supplemented with 5% vol/vol
fetal bovine serum (FBS; Gibco), 4.5 g/L D-glucose (Sigma),
0.2 mM L-glutamine (Gibco), 50 U/mL penicillin (Gibco), and
50 µg/mL streptomycin (Gibco). A fraction of the MNC was set aside
for cytospin slide preparation. Briefly, 20 000 MNCs/slide were
centrifuged in cytospin funnels at 500 rpm for 5 minutes using a
cytospin centrifuge (Shandon, Sewickley, PA).
Biotinylation of cells
Circulating cells.
Mature erythrocytes and circulating reticulocytes were biotinylated
with N-Hydroxysuccinimidobiotin (NHS-biotin; Pierce Chemical, Rockford,
IL). Biotin was dissolved at 10 mg/mL in dimethyl sulfoxide then
diluted into whole blood 1:1000 to make a concentration of 10 µg/mL.
After mixing in the dark for 5 minutes, cells were centrifuged and
washed twice in PBS (160 mM NaCl, 6.2 mM
KH2PO4, 25 mM Na2HPO4, 290 mOsm) and twice in hypotonic PBS, made by diluting PBS with deionized water to the desired osmolarity (155 mOsm).
Marrow cells.
Bone marrow MNCs were surface labeled with biotin by incubation in low
endotoxin phosphate-buffered saline (LE-PBS, Biowhittaker, Walkersville, MD) plus 5% FBS (Hyclone, Logan, UT) with water-soluble, reactive biotin (10 µg/mL EZ-Link sulfo NHS-LC-biotin, Pierce Chemical) for 5 minutes and stirred at room temperature in the dark.
Prior to use, the serum was incubated overnight with beads coated with
streptavidin (Dynal, M280 streptavidin beads) to remove biotin. Cells
were separated by centrifugation and washed 3 times in LE-PBS, then
suspended in filtered LE-PBS plus 5% FBS.
Mechanical measurements
In preparation for mechanical testing, thin glass fibers
(MO-SCI, Rolla, MO) were cemented (optical adhesive no. 68, Norland Products, New Brunswick, NJ) into the tapered tips of glass capillary tubes. A 3% gelatin solution (Sigma) was prepared in
Na2HPO4 (pH 7.2-7.4) and mixed vigorously with
5 mg NHS-biotin (Pierce Chemical) dissolved in 250 µL dimethyl
sulfoxide to form an emulsion. This solution was drawn into a glass
capillary and painted onto the tips of the fibers under a dissection
microscope. The gelatin was vapor fixed by suspending the painted fiber
in a test tube above concentrated formaldehyde (Sigma). On the day
before cells were tested, the fiber was mounted on the stage of an
inverted microscope (Nikon Diaphot, 60 × objective, with
monochromatic, 436-nm illumination, Nikon, Melville, NY) in a measuring
chamber filled with hypotonic PBS (~ 155 mOsm) containing BSA (3.0 mg/mL). Biotinylated RBCs were suspended at low concentration in 155 mOsm PBS plus BSA (3.0 mg/mL) and placed in the holding chamber
adjacent to the measuring chamber on the microscope stage.
Streptavidin-coated beads were introduced into the chamber and allowed
to adhere to the cells. Cell-bead pairs were selected and aspirated
into a large diameter (10 µm) transfer pipette. The pipette was
withdrawn from the holding chamber and introduced into the measuring
chamber and the cells were expelled. The transfer pipette was removed, and a calibration pipette, with an inside radius
(Rp) of 2.0 to 2.3 µm, was introduced. (The
RBCs suspended in the hypotonic buffer fit snugly into the pipette.)
Cell-bead pairs were manipulated into contact with the biotinylated
fiber. The cell was aspirated into the pipette and acted as a piston to
transfer the force of the aspiration to the fiber (Figure
1A). A series of known suction pressures
(
P) were applied to deflect the fiber. The force on the
fiber (f) was f = 
Rp2p. Typically, 3 to 5 different
calibration sequences were performed, each with a different cell, and
the average slope of the force deflection curves was used to
convert deflection to force. A typical calibration curve is shown in
Figure 1B. After calibration, the pipette was withdrawn, the chamber
was cleaned and filled with distilled water containing sodium azide (40 µM, Sigma) to retard bacterial growth, and the fluid level was
maintained via a connection to a fluid reservoir overnight.
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| Figure 1.
Calibration of a microcantilever.
(A). Video micrograph of the calibration of a microcantilever. The
biotinylated RBC adheres to the streptavidin-coated bead and serves as
a piston to transmit the force of the pipette suction pressure to the
fiber. The fiber deflection is measured as a function of the
suction pressure, and the force is calculated from the pressure as
described in the text. (B) An example of the force-deflection
relationship for a microcantilever. The force constant for this
cantilever was 5.6 pN/µm.
|
|
On the morning of the actual experiment, the chamber was flushed
thoroughly with physiologic saline and filled with LE-PBS plus 5% FBS
(vol/vol) for measurements. (The serum was preincubated with
streptavidin beads to remove biotin.) Biotinylated cells were suspended
at low density and placed in a holding chamber on the stage of the
microscope. Selected cells were transferred to the measurement chamber
and placed where they could be retrieved later. In experiments on
marrow cells, cells were selected based on their hemoglobin density and
morphology, and streptavidin beads were introduced into the chamber
after cells were transferred. In mature cell experiments, cells were
selected with beads already attached, and the cell-bead pairs were
transferred to the measurement chamber. When the appropriate
number of cells and beads had been moved into the measurement chamber,
the transfer pipette was replaced with a measurement pipette having an
inside diameter of 1.8 to 2.2 µm. Cells were attached to the
calibrated glass microcantilever via a streptavidin-coated bead, and a
portion of the cell was aspirated into the micropipette and withdrawn,
forming a tether (thin cylinder of bilayer) from the cell surface
(Figure 2). The deflection of the fiber
was used to determine the force required to form and maintain the
tether. In some cases, the dependence of the tethering force on
aspiration pressure was determined by applying a series of aspiration
pressures. After the formation of the tether and after each change in
pressure, the force was allowed to relax for 5 to 10 minutes to reach a
steady-state value.
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| Figure 2.
Tether formation.
Video micrographs show tether formation from a marrow reticulocyte
(left column) and a normoblast (right column). The top images show the
cell morphology and the resting position of the cantilever. As the
cells are withdrawn, the cantilever deflects, providing a measure of
the force (middle panels). When the force gets big enough, the surface
yields, and a strand of membrane is pulled out between the cell
and its attachment site on the bead. After a length of 10 to 30 µm is
reached, the cell is held stationary and the force on the tether is
allowed to relax to a steady value (bottom panels).
|
|
Statistics
Statistical significance was assessed by applying the
Student t test at a 95% confidence level.
Theory
For smooth-surfaced bilayer membrane capsules, the
mechanics of tether formation have been
delineated.14,16,17 For phospholipid vesicles, equilibrium
equations relating the pipette aspiration pressure, tether force, cell
dimensions, and material constants were obtained using either force
balance17 or energy methods.18 Energy methods
have been applied to obtain equilibrium relationships for tethers
formed from cells having linkages between the bilayer and the
underlying cytoskeleton.13,14 The dimensions of the system
are shown schematically in Figure 3. The
cell is held in a pipette with inside radius Rp
and the length of the cell projection into the pipette is
Lp. The pressure difference between the inside of the pipette and the surrounding buffer is P, and the
force on the tether is f. The length of the tether is
Lt and its radius is Rt.
The form of the energy function for the system is:
|
(1)
|
In this equation the first term corresponds to the energy
required to bend the membrane into a cylinder of radius
Rt. The energy is characterized by the bending
stiffness of membrane kc, which has units of
energy. The second and third terms represent the work of external
forces, and the fourth term corresponds to the energy needed to
separate the membrane bilayer from the underlying membrane skeleton and
associated proteins. This energy is represented by
Wsk, which has units of energy per unit area.
Contributions to the bending energy due to the relative stretching of
the adjacent leaflets of the membrane as a result of tether formation
are neglected because previous studies have shown that this energy is
not significant in RBCs for tethers up to 100 µm in length. (For more
detail on this energy term, see references 13 and 19.) To obtain
equations of equilibrium we take the variation of this function subject to the constraints that the membrane area and cell volume are constant.
Under these conditions, 2 independent equilibrium equations are obtained:
|
(2)
|
|
(3)
|
where Rc is the radial dimension of the
spherical portion of the cell outside the pipette. These are written as
approximations because terms on the order of the tether radius are
neglected in comparison with terms on the order of the cell radius.
These can be combined to eliminate the radius of the tether and obtain an expression for the tethering force as a function of the work of
separating bilayer from skeleton and the holding pressure in the
micropipette
![<IT>f<SUP>2</SUP></IT> = 8<IT>&pgr;<SUP>2</SUP> k<SUB>c</SUB></IT>[W<SUB><IT>sk</IT></SUB> + <IT>&tgr;</IT>]](/content/vol97/issue6/fulltext/1869/img004.gif)
|
(4)
|
where the membrane tension is given by:
|
(5)
|
Thus, the square of the tethering force is predicted to increase
linearly with the aspiration pressure, and the intercept of the line is
proportional to the energy cost of separating bilayer and skeleton
(Wsk).
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| Figure 3.
Schematic illustration of the tether formation
experiment showing the critical dimensions of the system.
The streptavidin-coated bead adheres to both the biotinylated
gelatin-coated fiber tip and the biotinylated cell surface. As the cell
is withdrawn a tether (cylindrical membrane strand) forms between the
cell and the bead. The deflection of the fiber provides a measure of
the force on the tether.
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|
The development of equation 4 involves 2 important assumptions about
the tether formation process. The first of these is that the system can
be treated as being in thermodynamic equilibrium. This condition is
verified by noting that the same equilibrium condition is reached
whether it is approached from increasing or decreasing tether length.
This is illustrated in Figure 4, in which
the tethering force and tether length are plotted as a function of time
for a typical tether formed from a marrow reticulocyte. When the tether
length is changed, the system is temporarily in disequilibrium, but
relaxes to an equilibrium state over a period of 5 to 10 minutes. The
source of this disequilibrium is not known, but it appears to be
related to the velocity of tether formation and the magnitude of the
change in tether length. When the difference between the instantaneous
tether force and the equilibrium tethering force is large, longer times
are required for the relaxation to equilibrium to occur.
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| Figure 4.
Time course of force relaxation after lengthening and
shortening of the tether.
Both the tether length (lower curve, filled symbols) and tether force
(upper curve, open symbols) are shown as functions of time. The
equilibrium force for this tether is between 34 and 35 pN. This value
is approached after the initial formation of the tether and again after
successive lengthening and shortening of the tether length. This
behavior is characteristic of a system in thermodynamic
equilibrium.
|
|
The second important assumption in developing equation 4 is that there
is no significant change in the elastic energy stored as a result of
the deformation of the cell when either the aspiration pressure was
changed, or the tether length was altered. For tether lengths that are
practical to achieve experimentally, the amount of material drawn into
the tether is too small to produce measurable deformations of the cell
body. Thus, the equilibrium tethering force is expected to be
independent of tether length, and this is confirmed experimentally. On
the other hand, elastic energy changes associated with cell deformation
resulting from changes in aspiration pressure are important to
consider, particularly in the present study because of the changes in
cytoskeletal organization and cell deformability that accompany the
maturation process. The effects of aspiration-induced cytoskeletal
deformation are manifested in an altered dependence of the tethering
force on aspiration pressure, which enters equation 4 via the term 
.
From the perspective of force balance, cytoskeletal rigidity results in
gradients in the membrane tension , such that the surface forces
generated by the aspiration pressure are not "felt" in regions far
from the pipette entrance. Thus, the degree to which the tethering
force depends on membrane tensions generated by pipette aspiration
() depends on the rigidity of the membrane-associated cytoskeleton,
or more specifically, how the elastic stiffness of the cytoskeleton
compares with the magnitude of the applied pressure. For mature RBCs,
the membrane rigidity is relatively small, and pressures can be applied
that make its contribution small. For less mature cells, the
cytoskeletal rigidity is larger, and this rigidity tends to reduce the
dependence of the tethering force on the aspiration pressure. In the
extreme case, the tethering force is independent of aspiration
pressure, and the contribution from the aspiration pressure does not
appear in the expression for force14:
f2 = 82 kc
Wsk.
This is an important consideration in the present study because the
changes in cytoskeletal organization during late-stage maturation have
significant effects on the relationship between tethering force and
aspiration pressure, as will be shown. Thus, differences in tethering
force for cells of different maturity may reflect not only differences
due to the intrinsic energy of association between bilayer and
skeleton, but also differences due to the degree to which cytoskeletal
rigidity influences the contribution of the holding pressure in the
pipette. To avoid these complications, direct comparisons of the
strength of the bilayer skeletal interactions should be made in the
limit as the aspiration pressure (contained in ) approaches zero.
| |
Results |
In an initial series of measurements, circulating reticulocytes
were tested and the equilibrium tethering force was compared to that of
control cells, that is, mature RBCs from unfractionated whole blood
(Figure 5). Tethers approximately 30 µm
in length were formed at a pipette holding pressure of approximately 98 Pa (1.0 cm H2O) and then the force was allowed to
"relax" for approximately 20 minutes to ensure that a steady value
was reached. (Note: The time required for the tether to reach an
apparent equilibrium depended on the conditions of the measurement. In
simple pulling experiments where the tether formation rate was high and
tethers were long, relaxation of the tethering force could be detected up to 15 minutes after pulling stopped. For experiments in which tethers were held at shorter, constant lengths and only the aspiration pressure was changed, relaxation of the tethering force occurred more
quickly, and the force appeared to reach equilibrium within 5 to 10 minutes of the change in pressure.) Forty-three circulating reticulocytes were tested and compared to tethering forces from 31 control cells. Although some cells in the reticulocyte population exhibited tethering forces below the range of forces observed for
control cells, the difference between the means of the 2 populations was not statistically significant. Subsequently, similar tests were
performed on hemoglobin-dense cells obtained directly from marrow.
(Cells were selected from the mononuclear fraction based on their
anucleate or normoblast-like appearance and on their dark color under
blue illumination.) The force of tether formation from these cells was
significantly less than the forces measured for mature RBCs. The mean
value for the steady-state tethering force for marrow cells, including
marrow reticulocytes and late-stage normoblasts, was 27 ± 9 pN
(mean ± SD, n = 37). This is significantly smaller than
tethering forces measured for mature red cells (54 ± 14 pN,
n = 31).13 In addition to these quantitative
differences, immature cells from marrow (unlike their mature
counterparts) frequently exhibited irregular contours and often changed
shape during testing. Interestingly, these changes in contour had very little effect, and in many cases no measurable effect at all, on the
magnitude of the tether force.
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| Figure 5.
Equilibrium tethering forces.
Histogram shows the distribution of equilibrium tethering forces for
mature cells (black bars), circulating reticulocytes (hatched bars),
marrow cells (including reticulocytes and normoblasts, gray bars), and
normoblasts only (gray, hatched bars). All tethers were formed at
membrane tensions ranging from 0.05 to 0.10 mN/m (aspiration pressures
ranging from 1.0 to 2.0 cm H2O). Although several
circulating reticulocytes exhibited tethering forces that fell below
the range of values for the mature cells, the difference between the
circulating reticulocyte population and the population of mature cells
was not statistically significant. However, tethering forces for
reticulocytes and normoblasts from marrow samples were significantly
lower than for mature cells.
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|
To quantify the changes in instability in terms of the separation
energy Wsk, the tethering force was measured as
a function of the pipette holding pressure. As expected (equation 4), a
linear relationship between the force squared and the apparent membrane tension was observed for both reticulocytes and mature RBCs (Figure 6A). The extrapolated intercept was used
to calculate the bilayer-skeletal separation energy
Wsk. Assuming that the bending stiffness of the
mature membranes and the reticulocytes was the same, and taking the
extrapolated values of ~ 1200 pN2 for mature cells
and ~300 pN2 for reticulocytes and normoblasts, we
estimate that membrane modifications during the last stages of
erythrocyte maturation result in a 4-fold increase in the energy
required to separate bilayer from skeleton from 19 µJ/m2
to 78 µJ/m2.
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| Figure 6.
Tethering forces as a function of pipette holding
pressure.
(A) The square of the equilibrium tethering force as a function of the
membrane tension for selected cells. Open symbols represent
measurements made on mature RBCs, dark symbols correspond to marrow
reticulocytes, and gray symbols correspond to normoblasts. Linear
regressions to each group of cells and 95% confidence intervals for
the fits are shown as solid lines amid the data. (B) Square of the
tethering force as a function of the membrane tension for a broader
representation of cells from marrow. A continuous range of behavior was
observed, from the force being independent of the membrane tension
induced by the aspiration pressure to a linear dependence of the force
squared on membrane tension. Intermediate behaviors shown here
probably reflect changes in cytoskeletal rigidity and a transition from
a highly wrinkled surface to the smooth contour of a nearly mature
reticulocyte. Each group of connected symbols represents a different
cell, all of which are marrow reticulocytes except the lowermost 4 curves (solid symbols: square, up-triangle, down-triangle and circle),
which are normoblasts.
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A range of behaviors was observed for the marrow population with regard
to the dependence of the tethering force on aspiration pressure (Figure
6B). Cells that were nonnucleated exhibited the greatest dependence of
force on pressure, and cells that had the largest nuclei and the most
ruffled surfaces exhibited almost no dependence. As discussed in the
previous section, this observation is consistent with the higher
mechanical stiffness of less mature cells.5-7 In addition,
the presence of a reserve of membrane created by surface wrinkling in
less mature cells makes it possible to form tethers from the cell
surface without drawing membrane from the pipette, and so without doing
work against the holding pressure. Thus, in addition to differences in
stability between marrow cells and mature cells, we observe a
progressive shift in behavior from that characteristic of a stiff,
ruffled membrane with local reservoirs of membrane surface area from
which tethers can be formed, to behavior characteristic of smooth
membranes, in which cells are sufficiently deformable that the excess
membrane area can be drawn completely into the micropipette, creating a smooth membrane contour. Thus, the observed changes in the dependence of the tethering force on aspiration pressure are consistent with the
decrease in cytoskeletal rigidity that is known to occur during late-stage erythroid maturation.
| |
Discussion |
A great deal is known about the composition and organization of
the membrane of mature RBCs, but surprisingly little is known about the
processes of protein synthesis and assembly that lead to the formation
of the mature cell. This is in part due to the difficulty of obtaining
immature RBCs in sufficient quantity and uniformity to perform
biochemical studies. RBCs are notoriously difficult to obtain in
culture, and marrow samples include cells of multiple lineages at
diverse stages of maturation. Consequently, most prior studies have
relied on transformed, erythroleukemic cells as a model system for
studying patterns of protein synthesis during erythropoiesis. Even in
this case, however, it is rare to bring erythroid precursors to the
fully mature form, and furthermore, interpretation of these studies may
be problematic because of alterations to the natural maturation
sequence that could result from transformation. Finally there is the
additional consideration that biochemical and ultrastructural studies
reveal information about structure and composition, but not about
function, and so, although there are a few studies that provide
information about the timing of the appearance of many of the membrane
skeletal proteins found in the mature cell, studies in which the
mechanical function of these assembled proteins is assessed are
extremely rare. An important and notable exception is the study
published by Chasis and coworkers7 examining changes in the
mechanical rigidity and stability of maturing reticulocytes and
assessing the importance of actin filaments and microtubules in
different aspects of the maturation process. The present findings
confirm many of the conclusions reached in that study and provide
additional quantitative measures of the changes in membrane stability
during the final stages of erythroid maturation.
The decrease in membrane stability that is documented here contrasts
with the increased rigidity of immature RBCs that has been documented
in a number of previous studies. In one of the first applications of
micropipette manipulation of RBCs, it was shown that it is
significantly more difficult to deform the surfaces of normoblasts and
reticulocytes than it is to deform the membranes of mature
cells.5 This finding has been confirmed subsequently by
others.6,7 Chasis and coworkers7 also found
evidence that immature RBCs are mechanically unstable, a conclusion
that is confirmed in the present study. This seemingly paradoxical result shows that there are important fundamental differences in the
mechanisms that account for resistance to membrane deformation, and
mechanisms that determine the stability of the membrane bilayer and the
strength of its association with the underlying
cytoskeleton.8
The changes in membrane stability observed in the present study reflect
the final stages of organization of the erythrocyte membrane skeleton
(or membrane-associated cytoskeleton), which, in the mature cell,
accounts for the elastic resistance of the membrane in extension
(surface shear deformation) and also acts to stabilize the membrane
bilayer against fragmentation.8,20,21 A number of
studies have documented the appearance and assembly of RBC membrane
proteins from the pronormoblast to the late normoblast. Decreased
turnover of spectrin and actin, and changes in the expression of
alternatively spliced forms of protein 4.1 are known to occur between
the early and late stages of normoblast maturation.4,22 In
the reticulocyte, reductions in lipid content and loss of surface receptors is known to occur during the final steps of maturation to the
mature cell.1,2 Chasis and coworkers have shown that microtubles and microfilaments play an active role in enucleation and
reticulocyte motility, respectively,7 but there is little information on the molecular reorganizations that occur in the membrane
skeleton and associated proteins during these final stages of
reticulocyte maturation. The findings of the present study confirm
conclusions reached previously that important changes in membrane
organization occur during the last 48 to 72 hours of reticulocyte
development, and demonstrate that these changes are essential for
establishing the stability and deformability of the mature cell.
For smooth vesicular membranes, the relationship between force and
aspiration pressure can be used to calculate the bending stiffness of
bilayer membranes.16 The mechanical analysis used to
obtain the relationship for calculating kc
includes the assumption that the membrane tension is uniform over the
cell surface. This assumption is valid for pure phospholipid bilayers
and for mature RBCs under conditions in which the membrane tension
generated by the aspiration pressure is large in comparison with
variations in membrane tension arising from the elastic deformation of
the membrane skeleton. The observation that the dependence of tethering force on aspiration pressure varies widely among normoblasts and marrow
reticulocytes of different maturity demonstrates that this assumption
is not valid for these cells. Thus, the apparent membrane bending
stiffness for reticulocyte membranes calculated from the slope of the
data obtained in the present study (0.886 × 10
19
J) is unlikely to be a true reflection of the intrinsic bending stiffness of the membrane. The value of 1.55 × 1019
J for mature cells is also somewhat lower than what has been reported in previous studies. Based either on measurements of surface
buckling, or on tether formation studies in which a different method of
calculation was used, it was found that kc for
mature cells falls in the range from 1.8 to 2.5 × 1019
J.12,13,23 Thus, refinements in the analytical
framework that account for surface elasticity are clearly needed. Until such an analysis is completed, an alternative approach would be to
determine the bending stiffness for the cells by a method in which the
tether radius is calculated based on the displacement of material from
the micropipette as the tether is formed.13,24 Unfortunately, the irregular and dynamic shape of marrow reticulocytes makes this approach unreliable because it presumes that the cell surface contour has a stationary shape. Thus, the heterogeneity of
properties and complex shapes of marrow cells make it appear unlikely
that a satisfactory analytical framework can be developed for
calculating kc from measurements of tether
formation for these cells. This complicates the comparison of the
energy costs for bilayer skeletal separation for different types of
cells, because Wsk appears always as a product
with the bending stiffness kc. (See equation 4.)
Our inability to measure kc directly for these cells requires us to make comparisons between cell types assuming that
the bending stiffnesses of the membranes are similar. Thus, although
large differences in the bending stiffness of membranes of different
maturity are not expected, the reader should keep in mind that part of
the reported differences in Wsk may be
attributable to differences in the bending stiffness of the different membranes.
Two other membrane systems have been tested by tether formation.
In neuronal growth cone, the forces required to form tethers are on the
order of 5.0 to 10.0 pN,14 considerably smaller than what
is measured for even the relatively unstable RBC precursors we have
examined here. These forces, assuming a membrane bending stiffness of
2.0 × 1019 J, correspond to a separation
energy of only 4.0 µJ/m2.14 This value
approaches the behavior of a pure phospholipid bilayer. Neutrophils
show a much higher resistance to surface loss. The minimum tethering
force for neutrophils was measured at 45 pN, corresponding to a
separation energy of 128 µJ/m2,25 of similar
order to what we observe for mature RBCs.
Membrane instability could account for the difficulties that have been
encountered in trying to produce viable mature RBCs from marrow
precursors in culture. The instability that we have documented in
maturing RBCs indicates that there are mechanisms at work within the
marrow (eg, close-packing of cells) that stabilize membranes against
surface loss during maturation. Indeed, it is well known that
erythropoiesis in vivo occurs in discrete characteristic foci,
erythoblastic islands, consisting of one or more central macrophages
(CM) surrounded by a group of maturing erythrocytes.26 The
cytoplasmic folds and processes of the CM interdigitate extensively within the erythoblastic island, covering 75% to 80% of the surface of each erythroblast and frequently penetrating deeply into cytoplasmic indentations in these cells.27 It has been postulated that
the close apposition of cell surfaces within the erythoblastic island might serve to facilitate intercellular communication and transfer of
chemical agents between the cells, and there is evidence that the CM
may also facilitate erythroid enucleation and surface remodeling. The
results of the present study lead us to speculate that these close
intercellular contacts may also serve to stabilize the maturing membrane against premature loss of surface, thus enabling the RBC to
maintain a high ratio of surface to volume during late-stage maturation.
In conclusion, micromechanical studies of the properties of RBC
precursors document an instability in the membranes of immature cells
compared with their mature counterparts. Marrow reticulocytes exhibit a
resistance to surface area loss through bilayer skeletal separation
that is comparable to what is measured for less mature normoblasts. In
contrast, circulating reticulocytes exhibit membrane stability that is
indistinguishable from the mature cell. These findings reveal that
there are important changes in the organization of membrane skeletal
proteins after enucleation and that the stability of the mature cell is
attained only in the last days of cell maturation.
| |
Acknowledgments |
The authors thank Ms Donna Brooks for technical support and Ms
Donna Phillips for help in preparing the manuscript.
| |
Footnotes |
Submitted March 29, 2000; accepted November 6, 2000.
Supported by the U.S. Public Health Service under National
Institutes of Health grant no. PO1-HL18208. Additional support was
obtained from the National Science Foundation (BES-9631670) and the
National Aeronautic and Space Administration (NAG-8-1382). A. M. is grateful for a fellowship from the Alexander S. Onassis Foundation.
The publication costs of this
article were defrayed in part by
page charge payment. Therefore,
and solely to indicate this fact,
this article is hereby marked
"advertisement"
in accordance with 18 U.S.C.
section 1734.
Reprints: Richard E. Waugh, Department of Pharmacology and
Physiology, 601 Elmwood Ave, Box 711, Rochester, NY 14642-8711; email:
waugh{at}seas.rochester.edu.
| |
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Abstract
Introduction
